ANIMAL  MICROLOGY 


THE  UNIVERSITY  OP  CHICAGO  PRESS 
CHICAGO,  ILLINOIS 


THE  BAKER  &  TAYLOR  COMPANY 

NEW  YORK 

THE  CAMBRIDGE  UNIVERSITY  PRESS 

LONDON 

THE  MARUZEN-KABUSHIKI-KAISHA 

TOKYO,  OSAKA,  KYOTO,  FUKUOKA,  SENDAI 

THE  MISSION  BOOK  COMPANY 

SHANGHAI 


ANIMAL  MICROLOGY 

PRACTICAL  EXERCISES  IN  ZOOLOGICAL 
MICRO-TECHNIQUE 


BY 

MICHAEL  F.  GUYER,  PH.D, 

Professor  of  Zoology  in  the  University  of  Wisconsin 
President  (1916),  The  American  Microscopical  Society 


WITH  A  CHAPTER  ON  DRAWING  BY 

ELIZABETH  A.  SMITH,  PH.D. 

Instructor  in  Zoology  in  the  University  of  Wisconsin 


REVISED  EDITION 


THE  UNIVERSITY  OF  CHICAGO  PRESS 
CHICAGO,  ILLINOIS 


rV 

BlOLOoY 


COPYRIGHT  1906  AND  igiy  BY 
THE  UNIVERSITY  OF  CHICAGO 


All  Rights  Reserved 


Published  November  1906 
Second  Impression  November  1910 
Second  Edition  February  1917 
Second  Impression  April  1919 
Third  Impression  August  1921 
Fourth  Impression  August  1922 


Composed  and  Printed  By 

The  University  of  Chicago  Press 

Chicago,  Illinois,  U.S.  A 


PREFACE  TO  THE  FIRST  EDITION 

For  the  past  ten  years  it  has  been  a  part  of  the  writer's  duties 
bo  give  instruction  in  microscopical  technique,  and  it  has  seemed 
to  him  that  there  is  need  for  a  series  of  practical  exercises  which 
will  serve  to  guide  the  beginner  through  the  maze  of  present-day 
methods,  with  the  greatest  economy  of  time,  by  drilling  him  in  a 
few  which  are  thoroughly  fundamental  and  standard.  The  book 
is  intended  primarily  for  the  beginner  and  gives  more  attention  to 
the  details  of  procedure  than  to  discriminations  between  reagents 
or  the  review  of  special  processes.  The  student  is  told  what  to  do 
with  his  material,  step  by  step,  and  why  he  does  it;  at  what  stages 
he  is  likely  to  encounter  difficulties  and  how  to  avoid  them;  if  his 
preparation  is  defective,  what  the  probable  cause  is  and  the  remedy. 
In  short,  the  book  attempts  to  familiarize  the  student  with  the  little 
" tricks"  of  technique  which  are  commonly  left  out  of  books  on 
methods  but  which  mean  everything  in  securing  good  results. 

A  very  brief,  non-technical  account  of  the  principles  of  the  micro- 
scope is  inserted  (Appendix  A)  with  the  idea  of  giving  the  student 
just  enough  of  the  theoretical  side  of  microscopy  to  enable  him  to  get 
satisfactory  results  from  his  microscope.  The  microscope  is  so  ably 
treated  in  the  excellent  works  of  Gage  (The  Microscope)  and  Car- 
penter (The  Microscope  and  Its  Revelations)  that  the  writer  feels 
himself  absolved  from  any  further  responsibility  in  this  matter. 

The  aim  of  the  entire  book  is  to  be  practical :  to  omit  everything 
that  is  not  essential;  and,  above  all,  to  give  definite  statements 
about  things.  Appended  to  each  chapter  is  a  series  of  memoranda 
which  serve  to  supply  additional  information  that  is  more  or  less 
pertinent  without  obscuring  the  main  features  of  the  method  under 
consideration. 

In  Appendix  B  the  formulae  for  a  number  of  the  most  widely 
used  reagents  are  given  with  comments  upon  their  uses  and  manipu- 
lation. Following  this  (Appendix  C)  is  a  concise  table  of  a  large 


4945.30 


vi  Animal  Micrology 

number  of  tissues  and  organs  with  directions  for  properly  pre- 
paring them  for  microscopical  study. 

Inasmuch  as  every  experienced  worker  has  his  own  "best" 
method  for  the  preparation  of  almost  any  tissue,  it  is  manifestly 
impossible  to  give  all  "best  methods"  in  such  a  table.  The  writer 
believes,  however,  that  the  student  will  find  the  methods  recom- 
mended all  good  ones  which  will  yield  satisfactory  results. 

In  Appendix  D  some  directions  are  given  for  collecting  and 
preparing  material  for  an  elementary  course  in  zoology. 

It  is  hoped  that  the  volume  will  prove  of  use:  (1)  as  a  class 
textbook;  (2)  as  a  guide  to  the  independent  individual  worker 
(teacher,  physician,  college  or  medical  student,  or  novice);  (3)  as 
a  reference  book  for  teachers,  in  the  preparation  of  material  for 
courses  in  elementary  zoology,  histology,  or  embryology. 

In  the  matter  of  expressing  his  obligations  the  writer  is  at  a 
loss  to  know  just  what  to  do.  Many  of  the  methods  in  microscopi- 
cal technique  have  been  handed  down  tradition-wise  from  one  worker 
to  another  until  their  origin  is  unknown;  they  are  the  accumulated 
experiences  of  several  generations  of  workers.  Furthermore,  many 
points  have  been  absorbed,  as  it  were,  by  the  writer,  from  fellow- 
workers  in  the  Universities  of  Chicago,  Nebraska,  and  Cincinnati, 
respectively;  consequently  the  obligation  cannot  be  specifically 
expressed.  Where  the  name  of  the  originator  of  a  method  is  known, 
due  credit  has  been  given.  The  books  to  which  the  author  is  most 
heavily  indebted  are  the  volumes  of  Gage  and  Carpenter,  already 
mentioned,  Lee's  Microtomist' s  Vade-Mecum,  Whitman's  Methods 
in  Microscopical  Anatomy  and  Embryology,  Hardesty's  Neurological 
Technique,  Foster  and  Balfour's  Elements  of  Embryology,  Minot's 
Laboratory  Text-book  of  Embryology,  Huber's  translation  of  the 
Bohm-Davidoff  Text-book  of  Histology,  Stohr's  Text-book  of  Histology, 
Mallory  and  Wright's  Pathological  Technique,  Bausch's  Manipu- 
lation of  the  Microscope,  and  the  Journal  of  Applied  Microscopy. 
Grateful  acknowledgment  is  also  made  to  the  various  manufacturers 
of  microscopical  instruments  and  appliances  for  the  loan  of  most  of 
the  cuts  which  have  been  used  in  this  volume. 

M.  F.  G. 


PREFACE  TO  THE  SECOND  EDITION 

The  favorable  reception  accorded  the  first  edition  of  Animal 
Micrology  has  encouraged  the  author  to  believe  that  a  second  edition, 
incorporating  some  of  the  many  new  methods  which  have  appeared 
during  the  past  ten  years,  would  be  equally  welcome.  The  general 
plan  of  the  book  has  not  been  altered  (see  Preface  to  the  First  Edi- 
tion, on  a  preceding  page),  although  changes  have  been  made  on 
nearly  every  page,  many  sections  have  been  entirely  rewritten,  and 
two  new  chapters,  one  on  "Cytological  Methods,"  the  other  on 
"Drawing,"  have  been  added.  The  chapter  on  drawing  has  been 
prepared  by  Dr.  Elizabeth  A.  Smith. 

In  spite  of  a  determined  effort  to  limit  the  book  to  its  former  size, 
it  has  expanded  by  over  fifty  pages.  For  every  method  dropped 
there  seemed  to  be  a  host  of  good  new  ones  demanding  recognition. 
These  in  the  main,  however,  have  been  left  to  the  encyclopedia  and 
the  various  technical  books  and  journals  listed  at  the  end  of  the 
volume.  As  in  the  first  edition,  the  policy  has  been,  not  to  attempt 
to  give  all  "best"  methods,  but  rather  to  select  representative  good 
ones  which  have  proved  their  worth  by  satisfactory  tests  in  American 
laboratories. 

Whatever  merit  the  new  edition  may  prove  to  have  over  that  of 
the  earlier  one  is  due  in  no  small  measure  to  the  many  helpful  sugges- 
tions of  my  colleagues  in  other  colleges  and  universities.  I  am 
particularly  indebted  in  this  respect  to  Professors  C.  E.  McClung, 
R.  R.  Bensley,  H.  McE.  Knower,  F.  L.  Landacre,  F.  C.  Waite, 
B.  M.  Allen,  George  R.  La  Rue,  Edward  L.  Rice,  F.  D.  Barker, 
R.  M.  Strong,  and  H.  L.  Wieman,  and  to  Doctors  Elizabeth  A.  Smith 
and  C.  H.  Heuser. 

M.  F.  G. 


vii 


CONTENTS 

PAGE 

INTRODUCTORY 1 

Apparatus  and  Supplies  Required,  1 ;  General  Rules,  5 

CHAPTER  I.     PREPARATION  OF  REAGENTS 7 

Practical  Exercises,  7;  Memoranda,  13. 

CHAPTER  II.    GENERAL  STATEMENT  OF  METHODS 15 

Killing,  Fixing,  and  Hardening,  16;  Washing,  17;  Dehydrating,  18; 
Preserving,  19;  Staining,  19;  Clearing,  21;  Mounting,  22;  Imbedding, 
22;  Affixing  Sections,  23;  Decolorizing,  24;  Bleaching,  24;  Corrosion, 
25;  Decalcification  and  Desilicidation,  25;  Injection  Methods,  25; 
Isolation  of  Histological  Elements,  25;  Normal  or  Indifferent  Fluids  for 
Examining  Fresh  Tissues,  26;  General  Scheme  for  Mounting  Whole 
Objects  (In  Toto  Preparations)  or  Sections,  26. 

CHAPTER  III.    KILLING  AND  FIXING 27  - 

Cautions,  27;  Alcohol  Fixation,  28;  Fixing  witn  Zenker's  Fluid,  28; 
Fixing  with  Bouin's  Fluid,  29;  Formalin  as  a  Fixing  Reagent,  29; 
Memoranda,  30. 

CHAPTER  IV.    SIMPLE  SECTION  METHODS 33 

Free-Hand  Section  Cutting,  33;  Memoranda,  34. 

CHAPTER  V.    THE  PARAFFIN  METHOD:  INFILTRATION  AND  SECTIONING     .       36  ^ 
The  Method,  36;   Memoranda,  42;  Difficulties  Likely  to  Be  Encoun- 
tered in  Sectioning  in  Paraffin,  and  the  Probable  Remedy,  46. 

CHAPTER  VI.    THE  PARAFFIN  METHOD:  STAINING  AND  MOUNTING     .  48  ^ 

Staining  with  Hematoxylin,  48;  Double  Staining  in  Hematoxylin  and 
Eosin,  50;  Double  Staining  in  Cochineal  and  Lyons  Blue,  50;  Staining 
with  Heidenhain's  Iron-Hematoxylin,  51;  Iron-Hematoxylin  with 
Other  Stains,  52;  Staining  in  Bulk  before  Sectioning,  52;  Paraffin 
Method  for  Delicate  Objects,  53;  Euparal  as  a  Mounting-  and 
Preservation-Medium,  54;  Memoranda,  55. 

CHAPTER  VII.    THE  CELLOIDIN  METHOD 59 

The  Method,  59;  Staining  Celloidin  Sections  in  Hematoxylin  and 
Eosin,  62;  Memoranda,  62. 

CHAPTER  VIII.    THE  FREEZING  METHOD 67    ' 

The  Method,  67;  Memoranda,  69. 

CHAPTER  IX.    METALLIC  SUBSTANCES  FOR  COLOR  DIFFERENTIATION     .       71 
A  Golgi  Method  for  Nerve  Cells  and  Their  Ramifications,  71 ;  Memo- 
randa on  Golgi    Methods,  72;  Other   Silver-Nitrate  Methods,  74; 
Memoranda  on  Silver  Methods,  75;  Gold-Chloride  Method  for  Nerve 
Endings,  76. 

ix 


x  Animal  Micrology 

PAGE 

CHAPTER  X.     ISOLATION   OF  HISTOLOGICAL  ELEMENTS.     MINUTE   DIS- 
SECTIONS       77 

Dissociation  by  Means  of  Formaldehyde,  77;  Isolation  of  Muscle 
Fibers  by  Maceration  and  Teasing,  77;  Maceration  by  Means  of 
Hertwig's  Fluid,  78;  Mall's  Differential  Method  for  Reticulum,  79; 
Minute  Dissection  and  Mounting  of  Various  Parts  of  Insects,  79; 
Memoranda,  80. 

CHAPTER  XI.    TOOTH,  BONE,  AND  OTHER  HARD  OBJECTS 81 

Sectioning  Decalcified  Tooth,  81;  Sectioning  Decalcified  Bone,  81; 
Sectioning  Bone  by  Grinding,  82;  Memoranda,  82. 

CHAPTER  XII.    INJECTION  OP  BLOOD  AND  LYMPH  VESSELS 83 

Red  Injection  Mass,  83;  Blue  Mass,  84;  Yellow  Mass,  84;  Injecting 
with  a  Syringe,  84;  Micro-Injection  of  Embryonic  Vessels,  88;  Cor- 
rosion Methods,  90;  Memoranda,  86-92. 

v  CHAPTER  XIII.    OBJECTS  OF  GENERAL  INTEREST:  CELL-MAKING,  FLUID 

MOUNTS,  "!N  TOTO"  PREPARATIONS,  ETC 93 

Turning  Cells  93;  Mounting  in  Glycerin  (Water  Mites,  Transparent 
Larvae),  94;  Killing  and  Mounting  Hydra,  95;  Mounting  in  Glycerin- 
Jelly  (Small  Crustacea,  etc.),  96;  Mounting  in  Balsam  (Flat  Worms, 
Mosquito,  Gnat,  Aphid),  97;  Opaque  Mounts  (Beetles,  Wings  of 
Moths  and  Butterflies,  Head  of  Fly,  Foreleg  of  Dytiscus),  98;  Dry 
Mounts,  99;  Spolteholz  Method  of  Clearing  Total  Specimens,  99; 
Memoranda  (Including  Directions  for  Mounting  Other  Forms),  99-104. 

CHAPTER  XIV.    BLOOD 105 

Examination  of  Fresh  Blood,  105;  Effects  of  Reagents,  105;  To 
Demonstrate  Blood  Platelets,  105;  Stained  Preparation  of  Fibrin,  105; 
Crystals  of  Blood,  105;  Cover-Glass  Preparations  (Dry),  106;  Rapid 
Method,  107;  Enumeration  of  Blood  Corpuscles,  107;  Observation  of 
the  Blood  Current,  109;  Inflammation,  109;  Memoranda,  110-111. 

CHAPTER  XV.    BACTERIA 112 

Bacterial  Examination,  112;  Cover-Glass  Preparations  from  Fluid 
Media,  112;  Staining  and  Mounting,  113;  Bacteria  in  Tissues,  113; 
Methylen  Blue  Stain  for  Bacteria  in  Tissues,  114;  Gram's  Method  for 
Bacteria  in  Tissues,  114;  Hanging-Drop  Preparations,  115;  Memo- 
randa, 115-118. 

CHAPTER XVI.    SOME  EMBRYOLOGICAL  METHODS:   SECTIONS  AND  WHOLE 

MOUNTS  OF  FROG  AND  CHICK;  OTHER  FORMS 119 

The  Frog,  119;  Section  Method,  119;  Whole  Mounts,  120;  Memo- 
randa on  Amphibian  Material,  121;  The  Chick,  123;  General  Memo- 
randa on  Embryological  Methods  and  Materials,  including  In  Vitro 
Cultures,  126-138. 


Contents  xi 

PAGE 

CHAPTER  XVII.    SOME  CYTOLOGICAL  METHODS 139 

General  Remarks,  139;  Mitosis,  139;  Testis  of  Crayfish,  Sections, 
140,  Smears,  141;  Blastodisk  of  Whitefish,  141;  Testis  of  Necturus, 
141;  Somatic  Cells  of  Ambystoma,  142;  Living  Cells,  143;  Mito- 
chondria, 143;  Staining  of  Living  or  Fresh  Tissues,  146;  Tests  for 
Certain  Cellular  Structures,  147;  Allen's  B-15  Method,  149;  Photo- 
graphing Cellular  Structures,  150;  Accessory  Chromosomes,  151; 
Acetc-Carmine  Preparations,  151;  Protoplasmic  Currents,  151; 
Celloidin  instead  of  Paraffin,  151;  Urea  in  Fixing  Fluids,  152;  Drop- 
Method  of  Changing  Fluids,  152;  Cooling  Tissues,  153;  Tissues  of 
Young  Adults  Desirable,  153;  Dissection  of  Living  Cells,  153;  Estima- 
tion of  Carbon  Dioxide,  153;  Holmgren's  Canals,  153;  Euparal,  153. 

CHAPTER  XVIII.     RECONSTRUCTION  OP  OBJECTS  FROM  SECTIONS     .     .      .     154 
Reconstruction    in    Wax,    154;     Geometrical   Reconstruction,    156; 
Blotting  Paper  Method,  157;   Rolling  Drawings  into  the  Wax,  157; 
Photography  in  Reconstruction  Work,  157;  Cutting  out  Wax  Plates, 
157. 

CHAPTER  XIX.     DRAWING 159 

Materials  for  Class  Work,  159;  Methods  of  Representation,  160; 
Outline,  160;  Depth,  160;  Ink  Drawings,  161;  Pencil  Drawings,  161; 
Wash  Drawings,  162;  Size  and  Arrangement  of  Drawings,  163;  Label- 
ing, 164;  Modes  of  Representation  for  Special  Courses,  164;  Embry- 
ology, 165;  Histology,  166;  Cytology,  166;  Drawings  f or  Publication, 
167;  Materials  for  Manuscript  Drawings,  167;  Camera  Lucida,  168, 
Reduction,  168;  Line  Process,  168;  Half-tone,  170;  Wash  and  Combi- 
nation Drawings  for  Reproduction,  170;  Lithography,  171;  Arrange- 
ment for  Reductions,  172;  Lettering,  172. 

APPENDIX  A.  THE  MICROSCOPE  AND  ITS  OPTICAL  PRINCIPLES  ....  175 
Optical  Principles,  175;  Images,  177;  The  Simple  Microscope,  178; 
The  Compound  Microscope,  179;  Defects  in  the  Image,  181;  Nomen- 
clature or  Rating  of  Objectives  and  Oculars,  183;  Some  Common 
Microscopical  Terms  and  Appliances  (Alphabetically  Arranged),  187; 
Manipulation  of  the  Compound  Microscope,  203. 

APPENDIX  B.    SOME  STANDARD  REAGENTS  AND  THEIR  USES     ....     207 
Fixing  and  Hardening  Agents,  207;  Stains,  218;  Normal  or  Indiffer- 
ent Fluids,  236;  Dissociating  Fluids,  237;  Decalcifying  Fluids,  238. 

APPENDIX  C.     TABLE  OF  TISSUES  AND  ORGANS  WITH  METHODS  OF  PREPA- 
RATION         ...     .  »  .     .      .     240 

APPENDIX    D.    PREPARATION    OF    MICROSCOPICAL    MATERIAL    FOR    A 

GENERAL  COURSE  IN  ZOOLOGY 259 

APPENDIX  E.    TABLE  OF  EQUIVALENT  WEIGHTS  AND  MEASURES     .      .      .     274 

APPENDIX  F.    REFERENCES 276 

INDEX       ...  279 


INTRODUCTORY 
APPARATUS  AND  SUPPLIES  REQUIRED 

The  student  should  provide  himself  with  the  following  supplies: 

One  half -gross  box  best  grade  glass  slides,  standard  size  (25X75  mm.). 
One-half  ounce,  18mm.  or  f  in.,  round  cover-glasses,  medium  thickness, 

(0.18mm.). 

Thirty  25X50  mm.  cover-glasses,  medium  thickness. 
Two  or  three  Pillsbury  slide  boxes  (Fig.  1). 
One  box  of  labels  for  slides. 
Three  to  six  camel's  hair  brushes  (Fig.  2). 
Six  pipettes  (Fig.  3). 
One  set  of  dissecting  instruments  as  follows: 

One  large  scalpel  or  cartilage  knife  (Fig.  4). 

One  small  scalpel  (Fig.  5). 

Two  needles  (Fig.  6). 

One  fine  straight  scissors  (Fig.  7). 

One  fine  straight  dissecting  forceps,  file-cut  points  (Fig.  8). 

One  blow-pipe  (Fig.  9). 

One  section  lifter  (Fig.  10). 
To  which  may  well  be  added: 

One  heavy  scissors  (Fig.  11). 

One  curved  scissors  (Fig.  12). 

One  heavy  forceps  (Fig.  13). 

One  fine  forceps,  curved  tips  (Fig.  14). 
One  horn  spoon. 

One  desk  memorandum  calendar. 
Blank  cards  (about  75X100  mm.)  for  keeping  records  of  experiments. 

The  kind  of  card  used  for  library  card  catalogue  will  do. 
One  section  razor  (Fig.  15). 
A  piece  of  moderately  heavy  copper  wire  with  one  end  hammered  out  to  a 

width  of  7  to  10  mm. 
Towels. 

A  glass-marking  pencil  (wax)  or  writing  diamond  will  be  found  useful. 
Coarse  carborundum  "engraver's  pencil  points,"  which  may  be  purchased 
for  seventy-five  cents  a  dozen,  are  very  satisfactory  for  marking 
3,  according  to  Professor  C.  E.  McClung. 
1 


Animal  Micrology 


FIG.  2 


FIG.  3 


FIG.  6 


FIG.  8  Fia.  9       FIG.  10 


Introductory 


FIG.  12  FIG.  13         FIG.  14 


FIG.  18 


^ 


FIG.  17 


FIG.  19 


FIG.  20  FIG.  21  FIG.  22 


FIG.  23 


4  Animal  Micrology 

Apparatus  ordinarily  supplied  by  the  laboratory: 

Desk  with  drawers. 

Locker  for  microscope. 

Compound  microscope  and  accessories  (Appendix  A). 

Dissecting  microscope  (Fig.  66). 

Microtomes  (Figs.  27,  28,  29,  32,  33). 

Paraffin  oven  (Figs.  24,  25,  26). 

Tall  stenders  (about  85mm.  deep).     Each  student  should  have  at  least 

eight  (Fig.  16). 
Coplin  staining  jars  (Fig.  17).    Tall  stenders  may  be  used  instead.    About 

eight  are  needed  for  each  student. 
Flat  stenders  (Fig.  18) ;  half  a  dozen  for  each  student. 
Syracuse  watch-glasses  (Fig.  19) ;  eight  to  each  student. 
Balsam  bottle  (Fig.  20). 

Graduated  cylinders  for  measuring  liquids  (Fig.  21). 
Wash-bottle  (Fig.  22). 
CeUoidin  bottle  (Fig.  23). 
Turntable  (Fig.  36). 
Injecting  apparatus. 
Reagent  bottles  and  vials. 
Other  apparatus  and  supplies  such  as  bone-forceps,  bone-saws,  glass  tubing, 

glass  rods,  beakers,  burners,  filter-paper,  funnels,  evaporating-dishes, 

sand  bath,  dropping-bottles,  balances,  mortar  and  pestle,  etc. 
For  apparatus  or  supplies  not  listed  in  this  book  the  student  is  referred 
to  the  illustrated  catalogues  of  dealers  and  manufacturers  such  as:  The 
Bausch  &  Lomb  Optical  Co.,  Rochester,  N.Y.;  The  Ernst  Leitz  Optical 
Works,  Wetzlar,  Germany  (American  branch,  30  E.  18th  St.,  New  York 
City);  The  Spencer  Lens  Co.,  Buffalo,  N.Y.;  Carl  Zeiss  Optical  Works, 
Jena,  Germany;  R.  &  J.  Beck,  68,  Cornhill,  London;  The  Kny-Scherer  Co., 
New  York  City;  Eimer  &  Amend,  New  York  City;  Whitall,  Tatum  &  Co. 
(especially  for  glassware),  New  York  City. 


IMPORTANT  GENERAL  RULES 

1.  Keep  everything  clean! 

2.  Have  a  definite  place  in  your  desk  for  each  piece  of  appa- 
ratus and  arrange  reagents  in  order  on  top  of  it. 

3.  Use-  cards  for   keeping   records   of   materials.     Each   card 
should  have  a  number  corresponding  to  that  of  each  special  object 
or  piece  of  tissue,  and  should  show  the  name  of  the  preparation, 
date,  reagents  used,  time  left  in  each  reagent — in  short,  all  data 
concerning  the  manipulation  of  the  material. 

4.  Jot  down  in  a  blank  calendar  the  various  things  to  be  done 
at  future  dates,  such  as  changing  of  reagent  on  tissues,  etc.,  and  then 
go  over  this  memorandum  carefully  each  day  when  you  first  come 
into  the  laboratory. 

5.  Use  only  clean  vessels  in  preparing  reagents,  and  clean  up 
all  glas^Tvare  while  it  is  yet  moist. 

6.  Reserve  and  mark  a  separate  pipette  for  each  of  the  chief 
reagents  (absolute  alcohol,  oils,  acids,  etc.). 

7.  In  making  up  solutions,  1  gram  of  a  salt  in  100  c.c.  of  liquid 
is  reckoned  ordinarily  as  a  1  per  cent  solution,  3  grams  as  a  3  per  cent 
solution,  etc.    But  if  solutions  are  to  be  of  10  per  cent  strength  or 
over,  it  is  better  to  weigh  out  the  dry  material  to  the  desired  percent- 
age and  then  add  enough  of  the  liquid  to  make  the  whole  weigh  100 
grams.    For  example,  to  make  a  25  per  cent  aqueous  solution  of 
caustic  potash,  add  25  grams  of  caustic  potash  to  75  c.c.  of  water.     A 
saturated  solution  contains  all  of  a  given  substance  that  the  liquid 
will  take  up.     When  a  solution  is  called  for  without  specifying  the 
solvent,  an  aqueous  solution  is  meant. 

8.  In  weighing  salts  always  first  put  paper  in  the  scale  pans  to 
protect  them. 

9.  In  making  solutions  or  mixtures  in  which  only  a  small  amount 
of  one  reagent  is  used,  after  mixing,  pour  back  some  of  the  mixture 
into  the  small  vessel  and  rinse  it  thoroughly  in  order  to  get  all  of  the 
original  contents  out. 

5 


6  Animal  Micrology 

10.  When  pouring  liquids  from  bottles  keep  the  label  of  the 
bottle  turned  toward  the  palm  of  the  hand.     Do  not  lay  down 
stoppers  but  hold  them  by  their  tops  between  the  knuckles. 

11.  Before  leaving  the  laboratory  put  away  your  instruments 
and  clean  and  put  in  its  place  whatever  laboratory  apparatus  you 
may  have  been  using. 

12.  All  solid  waste  materials,  acids,  stains,  etc.,  should  be  thrown 
into  stone  jars,  not  into  the  sink. 


CHAPTER  I 
PREPARATION  OF  REAGENTS 

The  following  reagents  should  be  prepared  by  each  student. 

1.  Grades  of  Alcohol. — To  obtain  a  given  percentage  of  alcohol 
through  dilution  of  a  higher  percentage  with  distilled  water,  sub- 
tract the  percentage  required  from  the  percentage  of  the  alcohol  to 
be  diluted;   the  difference  is  the  proportion  of  water  that  must  be 
added.     Thus,  if  35  is  the  percentage  required,  and  95  the  percentage 
to  be  diluted,  then  95—35=60;   hence,  60  parts  of  water  and  35 
parts  of  95  per  cent  alcohol  are  the  proportions  for  mixing. 

This  means  that  in  practice  one  needs  only  to  fill  the  graduated 
measuring  cylinder  to  the  same  number  as  the  percentage  required 
(e.g.,  35)  with  the  alcohol  to  be  diluted  (e.g.,  95)  and  then  fill  up 
to  the  percentage  of  the  latter  with  distilled  water.  In  this  way 
one  would  obtain  95  c.c.  of  alcohol  of  the  percentage  required,  if  the 
measuring  cylinder  is  graduated  in  cubic  centimeters. 

Prepare  about  250  c.c.  of  35,  50,  70,  and  83  per  cent  alcohols, 
respectively,  from  95  per  cent  alcohol  and  distilled  water.  The 
commercial  alcohol  used,  though  really  about  96  per  cent,  may 
be  figured  on  the  basis  of  95  per  cent. 

Owing  to  the  differences  in  the  specific  gravities  of  the  different 
percentages  of  alcohol,  the  foregoing  method  gives  only  approximate 
results;,  they  are  sufficiently  accurate,  however,  for  most  biological 
work. 

2.  Absolute  Alcohol. — It  is  customary  in  most  laboratories  to 
purchase  so-called  absolute  alcohol  specially  prepared  for  laboratory 
purposes.    Squibb's  absolute  alcohol  (99.8  per  cent)  is  commonly 
used.     Inasmuch  as  such  alcohol  is  an  expensive  reagent,  economy 
sometimes  necessitates  that  the  student  undertake  the  more  tedious 
process  of  making  his  own  absolute  alcohol.     Crystals  of  copper 
sulphate  are  heated  until  the  water  of  crystallization  is  driven  off 
and  the  sulphate  is  left  as  a  white  powder.     Such  anhydrous  sulphate 

7 


8  Animal  Micrology 

is  added  to  a  bottle  of  commercial  (96  per  cent)  alcohol.  The  water 
in  the  alcohol  immediately  unites  with  it,  turning  it  blue.  Anhy- 
drous sulphate  should  be  added  until  it  no  longer  turns  blue.  The 
alcohol  is  then  filtered  into  a  clean,  dry  bottle  which  must  have  a 
tight-fitting  cork  or  ground-glass  stopper.  It  is  well  to  smear  the 
glass  stopper  with  vaseline,  so  that  when  it  is  placed  in  the  bottle 
all  moisture  from  the  air  may  be  completely  excluded.  Any  labora- 
tory using  considerable  quantities  of  absolute  alcohol  should  have 
its  own  still. 

3.  Acid  Alcohol.— 

Alcohol  (70  per  cent) 99  c.c. 

Hydrochloric  acid  (pure) 1  c.c. 

For  sections  use  the  mixture  only  a  few  seconds  or  minutes. 
For  material  stained  in  bulk,  add  twice  as  much  70  per  cent  alcohol 
and  leave  the  object  in  it  until  sufficiently  decolorized  (2  to  24  hours). 

4.  Ether  and  Alcohol. — -Absolute  alcohol  and  sulphuric  ether 
equal  parts.     Quantity,  400  c.c.     Keep  the  ether  distant  from  all 
flames. 

5.  Normal  Saline. — Prepare  a  0.75  per  cent  solution  of  sodium 
chloride  in  distilled' w'Hter.     This  is  termed  a  normal  salt  solution 
because  it  is  a  solution  of  about  the  same  density  as  natural  lymph 
and  is  much  less  harmful  to  living  tissues  than  is  distilled  water. 
Quantity,  500  c.c.          «i 

6.  Formalin    (also   termed   formal,   formol,   formolose). — Com- 
mercial formalin  is  a  40  pjr  cent  solution  of  formaldehyde  in  water. 
A  4  per  cent  solution  of  formalin  would  be  made  by  taking  4  volumes 
of  commercial  formalin  and  96  volumes  of  water.     This  is,  howler, 
only  a  1 ,6  per  cent  solution  of  formaldehyde.     Make  a  10  per  cent 
solution  of  formalin.     Quantity,  250  c.c. 

7.  Zenker's  Fixing  Fluid.— 

Bichromate  of  potassium 2.5  grams 

Bichloride  of  mercury  (corrosive  sublimate)       5 . 0  grams 

Sodium  sulphate 1.0  gram 

Water 100     c.c. 

Glacial  acetic  acid 5     c.c. 


Preparation  of  Reagents  9 

Dissolve  the  bichromate  and  the  sublimate  in  the  water  with  the 
aid  of  heat.  Keep  the  acetic  acid  in  a  separate  bottle  until  the  fixing 
fluid  is  to  be  used,  as  it  will  produce  changes  in  the  chrome  salt  if 
added  at  once  and  allowed  to  stand. 

CAUTION. — In  handling  corrosive  sublimate  do  not  use  metal 
instruments  because  it  corrodes  metal.  Use  a  glass  or  horn  spatula. 

8.  Bouin's  Picro-Formol. — 

Picric  acid,  saturated  aqueous  solution 75  parts 

Formalin 25  parts 

Acetic  acid  (glacial) 5  parts 

One  gram  of  picric-acid  crystals  will  saturate  about  75  c.c.  of 
water. 

9.  MacCallum's  Macerating  Fluid. — 

Nitric  acid 1  part 

Glycerin 2  parts 

Water 2  parts 

10.  Decalcifying  Solution. — • 

Nitric  acid  (strong) 10  c.c. 

Alcohol  (70  per  cent) 90  c.c. 

11.  Alum-Cochineal. — 

Potassic  alum 6  grams 

Powdered  cochineal 6  grams 

Distilled  water 90  c.c. 

Boil  for  half  an  hour;  after  the  fluid  has  settled,  decant  the  super- 
nataVit  liquid,  add  more  water  to  it,  and  boil  it  down  until  only  90  c.c. 
of  the  decoction  remains.  •  Filter  when  cool,  and  add  a  bit  of  thymol 
or  a  little  salicylic  acid  to  prevent  the  growth  of  mold. 

12.  Delafield's  Hematoxylin. — Prepare  100  c.c.  of  a  saturated 
aqueous  solution  of  ammonia  alum.     Dissolve  1  gram  of  hematoxylin 
crystals  in  10  c.c.  of  absolute  alcohol,  and  add  it,  drop  by  drop,  to  the 
first  solution.     Expose  this  mixture  to  air  and  light  for  several  weeks 
(two  months  is  not  too  long)  to  "ripen."     (Ripening  consists  in  an 
oxidation  of  the  hematoxylin  to  form  hematein.    This  may  be  accom- 
plished at  once  with  some  degree  of  success  through  the  addition  of 
a  few  cubic  centimeters  of  a  neutralized  solution  of  peroxide  of 
hydrogen  or  other  powerful  oxidizing  agent.)     When  ripe,  filter  the 


10  Animal  Micrology 

solution  and  add  25  c.c.  of  glycerin  and  25  c.c.  of  methyl  alcohol  (see 
memorandum  1).  It  is  well  to  have  a  stock  solution  of  this  stain 
already  prepared  to  be  used  in  case  the  student's  preparation  is  not 
ready  in  time. 

Most  laboratories  keep  on  hand  a  stock  solution  of  hematoxylin 
made  by  dissolving  1  part  of  hematoxylin  crystals  in  10  parts  of 
absolute  alcohol.  In  the  course  of  several  months  or  a  year  this  solu- 
tion ripens  to  a  dark  wine-red  color.  It  may  then  be  used  in  making 
up  the  various  hematoxylin  solutions  and,  being  ripe,  will  stain  at 
once. 

NOTE. — At  this  point  the  student  should  begin  chap,  iii  in  order  that  no  time 
may  be  lost.  The  present  chapter  may  then  be  completed  while  the  tissues  are  becoming 
fixed  and  hardened. 

13.  Orange  G.— 

Orange  G  (Griibler's) .  . 1      gram 

Distilled  water 100      c.c. 

14.  Congo  Red.— 

Congo  red 0.5  gram 

Distilled  water 100     c.c. 

15.  Lyons  Blue.— 

Absolute  alcohol 100     c.c. 

Bleu  de  Lyon 0.3  gram 

16.  Eosin.— 

Eosin 0.5  gram 

Alcohol,  95  per  cent 100      c.c. 

17.  Iron-Hematoxylin  (Heidenhain's). — -Two  solutions  are  used. 
They  are  not  to  be  mixed. 

Solution  I: 

Ferric  alum  (clear  violet  crystals) 2.5  gram 

Distilled  water 100     c.c. 

Solution  II: 

Hematoxylin 0.5  gram 

Distilled  water. .  .   100      c.c. 


Preparation  of  E Agents  11 

The  hematoxylin  should  ripen  (see  I-eagent  12,  p.  9)  for  some 
three  or  four  weeks.  The  ferric  alum  of  the  histologist  is  always 
ammonio-ferric  sulphate. 


24.  —  The  Lillie  Water-Bath 


The  bath  consists  of  a  large  chamber  containing  a  series  of  drawers  of  equal  size 
250  mm.  long,  100  mm,  wide,  80  mm.  deep.  Each  drawer  has  copper  front  and  bottom; 
the  sides  and  back  are  perforated  zinc,  thus  securing  free  circulation  of  warm  air.  The 
drawers  are  separated  by  perforated  cross-partitions  and  run  on  slides  free  from  the 
lateral  supports,  thus  permitting  sufficient  circulation  of  warm  air  to  secure  equal  tem- 
perature in  the  top  and  bottom  of  the  bath.  Water  gauge  and  tubulatures  for  gas  regu- 
lator and  thermometer  are  provided.  This  bath  is  especially  adapted  to  class  work, 
since  each  student  may  carry  on  his  work  in  a  separate  drawer. 

18.  Canada  Balsam.  —  Dry  2  grams  of  Canada  balsam  on  a  sand 
bath  or  in  a  warm  chamber  until  it  becomes  hard  (1  to  2  hours  at 
65°  C.).  Do  not  overheat.  When  cool  add  enough  xylol  to  make 
a  very  thin,  syrupy  fluid.  Roll  a  sheet  of  paper  into  a  cone  to  serve 


12 


Micrology 


as  a  funnel,  and  filter  the  fRiid  through  absorbent  cotton.     Thicken 

the  solution  slightly  by  leaving  the 
cap  off  the  bottle  in  a  place  free 
from  dust,  and  allowing  some  of 
the  xylol  to  evaporate.  Or,  fill 
your  balsam  bottle  one-third  full  of 
the  liquid  xylol-balsam  now  on  the 
market  and  dilute  to  the  proper 
consistency. 

19.  Mayer's  Albumen  Fixative. 
— Beat  the  white  of  an  egg  with  an 
eggbeater  and  pour  it  into  a  tall 
cylinder.  Let  stand  until  the  air 
brings  all  suspended  matter  to  the 
top.  Skim  off  the  latter  and  to  the 
remainder  add  an  equal  volume  of 
glycerin,  and  a  bit  of  salicylate  of 
soda  (1  gram  to  50  c.c.)  or  thymol 
to  prevent  putrefaction. 

20.  Celloidin. — Soak  15  grams  of  dry  celloidin  overnight  in  just 
sufficient  absolute  alcohol  to  cover  it.     Then  dissolve  it  in  200  c.c.  of 
ether-alcohol  (see  reagent  4).     In  a 

second  bottle  prepare  a  thinner 
solution  by  taking  about  one-third 
of  the  original  and  diluting  it  with 
its  own  volume  of  ether-alcohol. 
The  solutions  are  best  kept  in 
bottles  with  glass  stoppers  and 
ground-glass  caps.  Label  the 
bottles  thick  and  thin  celloidin, 
respectively. 

21.  Paraffin. — In  one  of  the  cups 
of  a  warm  paraffin  oven  (Fig.  24, 
25,  or  26),  put  75  grams  of  paraffin, 
melting  at  about  50°  C.  (see  memo- 
randum  5,  p.   14).     The   bath  should  be  kept  at  a  temperature 


FIG.  25. — Simple  Water-Bath 
This  is  a  useful  bath  for  individual 
workers.  It  is  provided  with  imbed- 
ding-cups,  infiltration  vials,  a  shelf  for 
watch-glass  imbedding  or  for  warming 
instruments,  and  tubulatures  for  gas 
regulator  and  thermometer. 


PIG.  26. — Imbedding-Table 
There  should  be  two  rectangular 
boxes  (about  3X3X16  cm.)  to  contain 
paraffin.  When  in  use  the  boxes  are  so 
placed  on  the  imbedding-table  that  the 
paraffin  in  one  end  remains  melted;  in 
the  other,  solid.  Regulate  the  tem- 
perature by  placing  the  flame  at  the 
'proper  distance  under  the  acute  angle 
of  the  table.  It  is  best,  when  gas  is 
used,  always  to  turn  on  the  gas  com- 
pletely and  then  regulate  the  height  of 
the  flame  by  means  of  a  clamp  on  the 
rubber  tubing  which  conducts  gas  to 
the  burner. 


Preparation  of  Reagents  13 

of  some  two  degrees  above  the  melting-point  of  the  paraffin.  A 
supply  of  softer  and  of  harder  paraffin  (e.g.,  melting  at  43°  and  55°  C.) 
should  also  be  at  hand. 

Other  Reagents.— Provide  yourself  with  500  c.c.  of  distilled 
water,  200  c.c.  of  xylol,  25  c.c.  of  clove  oil,  25  c.c.  of  glacial  acetic 
acid,  50  c.c.  of  a  cedar-wood  oil,  25  c.c.  of  a  saturated  solution  of 
iodine  in  70  per  cent  alcohol,  75  c.c.  of  chloroform,  30  c.c.  of  glycerin, 
and  250  c.c.  of  absolute  alcohol  if  it  has  not  already  been  prepared. 
Keep  the  absolute  alcohol  and  the  xylol  carefully  corked  to  exclude 
moisture.  Before  measuring  out  any  of  these  reagents,  see  that 
both  the  graduate  and  the  bottle  are  perfectly  clean  and  dry. 

MEMORANDA 

1.  Ethyl  Alcohol  (grain  alcohol)  is  the  kind  commonly  used  in  histo- 
logical  laboratories.    Upon  presentation  of  the  proper  credentials  to  the 
internal  revenue  officers,  it  may  be  purchased  by  the  barrel  from  distillers, 
tax  free,  by  educational  institutions.    Such  commercial  alcohol  is  of  about 
96  per  cent  strength.    When  the  strength  is  unknown,  it  should  be  tested  by 
means  of  an  alcoholometer  (see  2,  below). 

Methyl  Alcohol  (called  also  wood  alcohol  or  wood  spirits)  is  cheaper 
than  ethyl  alcohol  in  case  the  latter  cannot  be  had  tax  free,  and  is  fairly 
satisfactory  in  most  cases.  It  is  poisonous  and  must  be  carefully  handled. 
It  is  of  about  90  per  cent  strength.  The  "methylated  spirits"  of  English 
microscopists  is  grain  alcohol  containing  10  per  cent  of  methyl  alcohol. 

Rectified  Spirit  is  a  91  per  cent  alcohol  (84  per  cent  in  England). 

2.  The  Alcoholometer  is  a  convenient  instrument  for  determining  the. 
strength  of  alcohol,  or  the  percentage  of  absolute  alcohol  in  a  spirituous 
mixture.     It  is  a  kind  of  hydrometer  with  a  scale  marked  to  indicate  the 
percentages  of  alcohol.     Different  strengths  of  alcohol  have  different  Specific 
gravities;  consequently  the  instrument  will  float  higher  or  lower  in  the  liquid 
according  to  the  percentage  of  alcohol  present.    The  number  on  the  scale 
just  at  the  surface  of  the  liquid  indicates  its  strength. 

3.  Rule  for  Dilution  of  a  given  strength  of  a  solution  with  a  lower  per- 
centage of  the  same  solution.     (For  where  the  diluent  is  water,  i.e.,  zero 
per  cent,  see  rule  under  reagent  1.)     Subtract  the  percentage  required  from 
the  percentage  of  the  solution  to  be  diluted;  also  subtract  the  percentage  of 
the  diluent  from  that  of  the  strength  required.    The  differences  are  the 
relative  proportions  of  the  diluent  and  the  solution  to  be  diluted  that  must 
be  used.     Thus,  to  prepare  a  35  per  cent  solution  from  95  and  20  per  cent 
solutions:    95-35=60;    35-20=15;    hence,   60  to  15  or  4  to  1  are  the 


14  Animal  Micrology 

proportions  desired.    That  is,  4  parts  of  the  20  per  cent  and  1  part  of  the 
95  per  cent  solution  must  be  used  to  obtain  a  35  per  cent  solution. 

4.  "To  Remove  Fixed  Stoppers,  take  the  bottle  in  the  left  hand  with  the 
thumb  applied  to  one  side  of  the  stopper,  then  tap  the  other  side  of  the 
stopper  with  some  heavy  instrument,  such  as  the  handle  of  a  pocket-knife, 
pressing  the  thumb  against  the  direction  of  the  tap.    Turn  the  bottle 
round,  gradually  tapping  until  the  stopper  loosens.    Should  this  device 
prove  of  no  avail  (which  is  very  rarely),  hold  the  neck  of  the  bottle  in  a 
spirit  flame,  and  quickly  withdraw  the  stopper  as  the  glass  of  the  neck 
expands.    This  is  a  somewhat  risky  procedure,  but  is  very  effectual  if  done 
smartly"  (Journal  of  Applied  Microscopy,  VI,  2116).    The  glass  of  the  neck 
may  be  more  safely  heated  by  looping  a  heavy  cord  about  it  and  sawing  the 
cord  back  and  forth  until  the  friction  warms  the  glass. 

5.  A  Simple  Paraffin  Bath,  recommended  by  several  workers,  may  be 
made  by  suspending  an  electric-light  bulb  (carbon-filament)  in  a  tumbler 
of  paraffin.    The  height  of  the  bulb  should  be  so  adjusted  that  some 
unmelted  paraffin  remains  at  the  bottom.    Tissues  will  thus  come  to  lie 
where  the  paraffin  is  j  ust  at  the  melting-point.    Professor  F.  C .  Waite  recom- 
mends putting  a  paper  cone  around  the  entire  apparatus  if  the  room  is  cold. 
McClendon,  in  the  Biological  Bulletin  (XV,  No.  1)  for  June,  1912,  explains 
how  to  make  a  convenient  concrete  paraffin  bath  for  individual  use  which 
is  inexpensive  and  effective. 

McClung  suspends  a  50-candle-power  helix-wound  carbon-filament 
lamp,  provided  with  a  lamp  shade,  over  a  beaker  of  paraffin.  The  heat 
from  the  lamp  is  sufficient  to  keep  the  top  of  the  paraffin  melted.  Rapid 
evaporation  of  the  dealcoholizing  agent  is  facilitated  by  such  an  arrangement 
and  overheating  is  avoided.  The  same  lamp  may  be  used  for  spreading 
the  paraffin  ribbon  on  slides,  and  for  drying. 


CHAPTER  II 
GENERAL  STATEMENT  OF  METHODS 

Each  of  the  reagents  which  has  been  prepared  is  used  for  one 
or  more  of  the  purposes  to  be  discussed  in  this  chapter. 

All  methods  of  preparation  in  microscopy  are  to  enable  us  to 
learn  more  of  the  structure  and  functions  of  objects  than  would 
otherwise  be  apparent.  We  endeavor  to  study  them  in  as  near 
their  natural  condition  as  possible.  While  the  study  of  living  or 
of  fresh  material  is  desirable  it  can  be  carried  on  only  to  a  very 
limited  extent.  Most  structures  of  the  animal  body,  though  opaque, 
must  be  examined  largely  by  transmitted  light,  hence  special  prepara- 
tion is  necessary  to  put  them  into  suitable  condition.  This  is 
accomplished — 

1.  By  cutting  them  into  thin  slices  (section  method). 

2.  By  separating  them  into  their  elements  (isolation) — 
a)  Mechanically  (teasing),  or 

6)  With  the  aid  of  fluids  which  remove  the  cement  substance 

(dissociation  or  maceration). 

In  most  instances  the  minute  structure  of  a  tissue  or  of  an  organ- 
ism can  be  studied  to  the  best  advantage  only  after  the  application 
of  certain  agents  which  serve  to  emphasize  the  various  structural 
elements.  A  tissue  so  prepared  is  an  artificial  product  in  that  it  is 
not  exactly  the  same  as  it  was  in  the  living  organism,  but  recent 
studies  of  protoplasm  in  the  living  condition  by  competent  investi- 
gators strengthen  the  belief  that  many  reagents  preserve  very 
faithfully  the  actual  structure  of  the  cell  contents.  The  liquid 
albuminoids  are  apparently  the  materials  which  suffer  the  greatest 
modifications.  Since  alterations  do  occur,  however,  it  is  clear  that 
in  our  interpretations  of  prepared  material  we  must  reckon  carefully 
both  with  the  original  nature  of  the  object  and  with  the  factors 
introduced  by  ourselves. 

15 


16  Animal  Micrology 

KILLING,  FIXING,  AND  HARDENING 

The  first  step  in  the  preparation  of  tissues  ordinarily  is  the 
employment  of  some  reagent  which  will  kill  the  tissues  and  fix 
their  various  components  in  the  characteristic  stages  of  their 
activities.  Such  material  may  then  be  preserved  indefinitely  for 
future  use. 

It  is  customary  to  discriminate  between  killing,  fixing,  and 
hardening,  although  the  same  reagent  may  fulfil  all  three  require- 
ments. Killing  refers  particularly  to  the  destruction  of  the  life  of 
the  tissue,  a  process  which  may  be  either  slow  or  instantaneous. 
In  slow  killing  it  is  usual  to  employ  narcotics  such  as  ether,  chloro- 
form, chloral  hydrate,  chloretone,  carbon  dioxide,  nicotin,  cocain,  or 
weak  alcohol.  Ice  is  also  used  sometimes.  Such  methods  are  of 
particular  value  with  highly  contractile  animals  which  are  desired 
in  the  extended  condition.  Such  forms  are  narcotized  completely 
or  until  they  are  unable  to  contract,  and  then  frequently  fixed  and 
hardened  in  other  or  stronger  fluids.  Where  practicable,  instantane- 
ous killing  and  fixing  is  preferable  because  tissues  have  then  no  time 
to  undergo  postmortem  changes.  The  same  fluid  ordinarily  is 
employed  for  killing  and  fixing. 

The  purpose  of  fixation  is— 

a)  To  preserve  the  actual  form  of  tissue  elements. 

6)  To  produce  optical  differences  in  structure,  or  so  to  affect 
the  tissues  that  such  differences  will  be  brought  out  through  sub- 
sequent treatment  with  stains  or  other  reagents. 

To  accomplish  this  the  fixing  agent  must  possess  the  following 
properties: 

1.  It  should  kill  the  tissue  so  quickly  that  few  structural  changes 
can  occur. 

2.  It  should  neither  shrink  nor  distend  the  tissue. 

3.  It  should  be  a  good  preservative;  that  is,  it  must  render  the 
tissue  elements  insoluble  and  prevent  postmortem  changes. 

4.  It  should  penetrate  all  parts  equally  well. 

5.  It  should  put  the  tissue  in  condition  to  take  stains  unless  it 
of  itself  produces  sufficient  optical  differences  in  the  various  parts  of 
the  tissue. 


General  Statement  of  Methods  17 

No  ideal  single  reagent  has  been  discovered  which  meets  all 
of  these  requirements,  hence  it  is  customary  to  combine  two  or 
more  reagents  which  individually  possess  certain  of  these  desirable 
qualities.  All  of  the  best  fixing  reagents  are  mixtures.  For  example, 
acetic  acid  is  very  generally  used  in  fixing  mixtures  because  it  pene- 
trates well,  produces  good  optical  differentiation,  and  counteracts  the 
tendency  of  some  reagents  (e.g.,  corrosive  sublimate)  to  shrink  tissues. 
Again,  osmic  acid,  which  is  an  excellent  fixing  agent  for  very  small 
pieces  of  tissue,  penetrates  very  poorly;  consequently  for  most 
objects  it  must  be  mixed  with  reagents  which  penetrate  rapidly  and 
thoroughly. 

Some  fixing  agents  (corrosive  sublimate,  chromic  acid,  osmic 
acid,  etc.)  enter  into  chemical  combination  with  certain  of  the  tissue 
elements;  others  (alcohol,  picric  acid,  nitric  acid,  hot  water,  etc.) 
act  by  coagulating  or  precipitating  certain  constituents  of  tissues. 

The  chief  object  of  hardening  is  to  bring  tissues  to  the  proper 
consistency  for  cutting  sections.  The  process,  although  begun 
ordinarily  by  the  fixing  agent,  is  usually  completed  in  alcohol. 
Some  objects  are  not  sufficiently  hardened  until  they  have  remained 
in  alcohol  for  many  hours,  or  even  days.  As  a  rule,  tissues  should 
remain  in  alcohol  of  at  least  70  per  cent  strength  for  a  minimum 
of  24  hours  after  the  preliminary  operations  of  fixing,  washing,  etc., 
before  they  are  subjected  to  further  treatment. 

WASHING 

Fixing  agents  ordinarily,  with  the  exceptions  of  alcohol  and 
formalin,  must  be  washed  out  thoroughly  or  they  are  likely  to  inter- 
fere with  subsequent  processes.  Aqueous  solutions  are  washed  out 
usually  in  water  or  a  low  percentage  of  alcohol;  alcoholic  solutions, 
with  alcohol  of  about  the  same  strength  as  that  of  the  fixing  agent. 
Washing  usually  requires  from  10  to  24  hours,  with  several  changes 
of  the  liquid.  If  water  is  the  washing  agent,  it  is  best  where  prac- 
ticable to  use  running  water. 

Chromic  acid  and  its  compounds  should  be  washed  out  in  run- 
ning water.  This  should  be  done  in  the  dark  in  order  that  pre- 
cipitation may  be  avoided. 


18  Animal  Micrology 

Picric  acid,  or  solutions  containing  it  (except  picro-formalin 
mixtures),  must  be  washed  in  strong  alcohol  (70  per  cent),  never  in 
water,  because  the  latter  seems  to  undo  the  work  of  fixation. 

Corrosive  sublimate  and  mixtures  containing  it  are  washed  out 
in  water  or  alcohol.  A  little  tincture  of  iodine  should  be  added 
to  the  wash  from  time  to  time  to  insure  the  removal  of  all  corrosive- 
sublimate  crystals.  Sufficient  iodine  has  been  added  when  it  no 
longer  loses  its  reddish  color  after  being  in  contact  with  the  prepara- 
tion for  a  short  time. 

Osmic  acid  and  mixtures  containing  it  should  be  washed  in  running 
water. 

DEHYDRATING 

While  under  certain  circumstances  objects  may  be  mounted  in 
aqueous  media  for  examination,  in  the  majority  of  cases,  especially 
where  the  preparation  is  to  be  a  permanent  one,  it  has  been  found 
best  to  remove  all  water  from  the  tissues,  that  is,  to  dehydrate  them. 
This  renders  preservation  more  certain,  and  it  is  a  necessity,  more- 
over, if  the  object  is  to  be  imbedded  later  in  paraffin  or  celloidin,  for 
neither  of  these  substances  is  miscible  with  water.  Because  of  its 
strong  affinity  for  water  and  the  ease  with  which  it  may  be  manipu- 
lated, alcohol  has  come  to  be  used  universally  for  this  purpose.  It 
completes  the  process  of  hardening  at  the  same  time.  The  dehy- 
dration must  be  gradual.  In  tissues  transferred  from  water  or 
aqueous  solutions  directly  to  strong  alcohol  (or  vice  versa)  violent 
diffusion  currents  are  set  up  which  produce  serious  distortion  of  the 
tissue  elements.  For  this  reason  a  series  of  alcohols  of  gradually 
increasing  strength  (e.g.,  35-50-70-83-95  per  cent)  is  used.  The 
more  delicate  the  object,  the  closer  should  be  the  grades  of  alcohol. 

Professor  C.  E.  McClung  recommends  a  "drop"  method  for  all 
purposes.  If,  for  example,  an  object  in  water  is  to  be  carried  up 
into  the  higher  alcohols,  he  places  a  vessel  containing  95  per  cent 
alcohol  and  the  vessel  containing  the  object  in  water  under  a  bell-jar. 
The  vessel  containing  alcohol  is  raised  above  the  level  of  the  other 
vessel  and  a  string  or  a  capillary  siphon  is  set  to  carrying  over  the 
alcohol  drop  by  drop  into  the  water.  The  amount  of  95  per  cent 
alcohol  must  be  apportioned  to  the  amount  of  water  so  that  the  final 


General  Statement  of  Methods  19 

desired  strength  will  be  reached  by  the  time  the  alcohol  has  all  passed 
over  into  the  water.  The  vessel  containing  the  specimen  should  be 
agitated  frequently  to  secure  thorough  mixture.  Obviously  other 
fluids  may  be  changed  by  the  same  method.  For  a  more  elaborate 
form  of  drop  apparatus  see  memorandum  6,  p.  152. 

PRESERVING 

After  fixing  and  washing,  the  process  of  dehydration  is  begun 
ordinarily  and  tissues  are  carried  as  far  as  70  per  cent  alcohol.  It 
is  customary  to  leave  them  in  alcohol  of  from  70  to  83  per  cent 
strength  until  they  are  needed.  They  may  remain  here  indefinitely. 
If  they  are  to  be  preserved  for  a  long  time  (for  months),  however, 
it  is  better  to  keep  them  in  a  mixture  of  equal  parts  of  glycerin, 
distilled  water,  and  strong  (commercial)  alcohol. 

STAINING 

A  few  fixing  agents  produce  sufficient  optical  differentiation  in 
tissues,  but  as  a  rule  this  must  be  accomplished  through  the  addi- 
tion of  certain  stains.  Most  of  the  stains  used  have  more  or  less  of  a 
selective  action;  that  is,  they  pick  out  certain  elements  of  the  tissue, 
and  thus  enable  one  to  see  details  of  structure  that  would  otherwise 
be  invisible.  Their  action,  however,  depends  largely  upon  the  nature 
of  the  fixing  agent  which  has  previously  been  used.  The  secret  of 
good  staining,  indeed,  lies  largely  in  proper  fixation. 

There  are  large  numbers  of  stains  of  very  different  chemical 
constitution  (acid,  neutral,  and  alkali),  and  they  may  act  in  very 
different  ways  upon  the  material  to  be  stained.  For  example,  some 
show  affinity  only  for  certain  elements  of  the  nucleus,  others  for  the 
cytoplasm  of  cells,  and  some  are  present  in  tissues  only  physically 
as  deposits,  while  others  enter  into  chemical  combination  with  cer- 
tain of  the  cell  constituents.  A  few,  such  as  borax-carmine,  are 
general  stains,  and  affect  to  a  greater  or  less  degree  practically  all 
the  tissue  elements. 

It  is  not  the  purpose  of  the  present  book  to  enter  into  a 
prolonged  discussion  of  the  theory  of  staining  or  to  undertake  a 
description  and  classification  of  stains.  For  this  the  reader  is 


20  Animal  Micrology 

referred  to  the  excellent  compendium  of  Lee  (The  Microtomist's 
Vade-Mecum). 

The  stains  of  widest  application  are  (1)  the  Carmines,  (2)  the 
Hematoxylins,  (3)  the  Anilins,  and  (4)  Metallic  substances. 

Carmine  is  a  brilliant  scarlet  or  purplish  coloring  matter  made 
from  the  bodies  of  the  cochineal  and  kermes  scale  insects.  The 
carmine  stains,  including  cochineal,  have  been  largely  used  in  the 
past  for  all  kinds  of  work,  but  at  present  they  are  used  more  par- 
ticularly for  staining  objects  in  bulk  before  sectioning,  or  objects 
which  are  not  to  be  sectioned.  They  are  easy  to  use,  and  will 
follow  almost  any  fixing  agent.  In  case  of  overstaining,  weak 
hydrochloric  acid  (0 . 1  to  1  per  cent)  is  used  to  decolorize  the  tissues. 
For  formulae  see  Appendix  B,  p.  220. 

Hematoxylin  is  a  compound  containing  the  coloring  matter  of 
logwood.  The  hematoxylins  follow  well  almost  any  of  the  fixing 
agents;  they  are  especially  recommended  after  fluids  containing 
chromic  acid  or  its  salts.  According  to  Mayer,  the  active  agent 
in  these  stains  is  a  compound  of  hematein  with  alumina.  This 
blue-colored  solution  is  precipitated  in  tissues,  particularly  in  nuclei, 
by  certain  organic  and  inorganic  salts,  such  as  phosphates.  The 
hematein  is  produced  by  the  oxidation  of  hematoxylin.  The  so- 
called  "  ripening "  is  simply  this  change,  which  is  brought  about 
by  exposing  the  hematoxylin  solution  to  air.  If  the  pure  hematein 
is  used  in  making  the  stain,  therefore,  the  latter  will  be  ready  for 
use  immediately,  because  it  need  not  undergo  the  ripening  process 
(se'e  remarks  under  12,  p.  9).  For  formulae  see  Appendix  B,  p.  225. 

Anilin  is  a  colorless  coal-tar  derivative,  and  is  the  base  from 
which  many  of  the  numerous  coal-tar  dyes  are  made.  The  anilins 
are  brilliant  stains  of  all  colors.  They  are  used  almost  exclusively 
for  staining  sections  or  thin  membranes,  and  are  of  great  service 
to  the  microscopist,  although,  as  a  rule,  they  fade  in  time. 

The  basic  anilin  stains,  such  as  methyl  green,  methyl  violet, 
gentian  violet,  methylen  blue,  safranin,  Bismarck  brown,  toluidin 
blue,  and  thionin,  are  usually  nuclear  stains.  On  the  other  hand, 
the  acid  anilin  stains,  such  as  acid  fuchsin,  eosin,  erythrosin,  light 
green,  orange  G,  bleu  de  Lyon,  nigrosin,  benzopurpurin,  and  aurantia, 


General  Statement  of  Methods  21 

are  ranked  as  cytoplasmic  stains.  Some  of  these  stains  must  be 
made  up  fresh  every  two  or  three  months,  as  they  frequently  spoil 
if  kept  much  longer. 

The  metallic  substances  used  for  color  differentiation  operate 
principally  as  impregnations  rather  than  as  stains.  The  coloring 
matter  is  held  physically  as  a  precipitate  or  reduction  product  in 
certain  of  the  tissue  elements.  The  commonest  reagents  of  this 
class  in  use  are  silver  nitrate  and  gold  chloride. 

The  different  tissue  elements  frequently  show  affinity  for  different 
stains;  consequently  it  is  a  common  practice  to  use  more  than  one 
stain.  Very  decided  contrasts  may  thus  be  produced,  such  as  red 
and  blue,  red  and  green,  green  and  orange,  etc.  It  is  not  uncommon, 
in  fact,  to  have  triple  and  even  multiple  staining.  In  such  staining 
the  stains  are  sometimes  applied  consecutively;  in  other  cases,  at 
different  points  in  the  process  of  general  manipulation.  Sometimes 
all  the  stains  may  be  mixed  together,  so  that  immersion  of  the  sections 
in  one  liquid  is  all  that  is  required  for  double  or  multiple  staining. 

A  general  rule  in  staining,  especially  for  entire  or  bulky  objects, 
is  that  the  specimen  should  be  transferred  to  the  stain  from  a  reagent 
in  which  the  percentage  of  water  is  approximately  the  same  as  that 
of  the  stain.  The  same  is  true  when  the  object  is  removed  from  the 
stain.  For  example,  if  the  stain  to  be  used  is  an  aqueous  solution, 
the  object  should  enter  it  from  an  aqueous  solution;  if  the  stain  is 
made  up  in  95  per  cent  alcohol,  the  object  should  enter  from  95  per 
cent  alcohol,  etc.  For  reasons  see  "Dehydrating." 

CLEARING 

In  the  vast  majority  of  cases  tissues  are  too  opaque  for  satis- 
factory examination  until  they  have  been  treated  with  certain 
clarifying  reagents  or  clearers  which  render  them  more  transparent. 

Such  reagents  as  glycerin,  glycerin-jelly,  etc.,  are  used  when  the 
object  is  to  be  cleared,  without  alcoholic  dehydration,  directly  from 
water.  Usually,  for  permanent  preparations,  the  alcoholic  dehydra- 
tion method  is  employed  and  it  then  becomes  necessary  to  use  a 
clarifying  reagent  which  will  replace  the  alcohol  and  facilitate  the 
penetration  of  the  final  mounting-medium  (balsam  or  damar). 


22  Animal  Micrology 

Perhaps  the  most  useful  and  rapid  clearer  is  xylol.  Xylol,  how- 
ever, is  very  sensitive  to  moisture,  and  if  the  preparation  has  not 
been  thoroughly  dehydrated  the  final  mount  will  appear  milky. 
Cedar-wood  oil,  though  somewhat  slower  than  xylol,  is  one  of  the 
best  clearers.  It  is  also  one  of  the  safest,  because  tissues  may  be 
left  in  it  indefinitely.  Other  good  clearers  after  alcohol  are  oil  of 
origanum,  sandal-wood  oil,  oil  of  cloves,  cassia  oil  (cinnamic  alde- 
hyde), toluol,  oil  of  bergamot,  anilin  oil  (for  watery  specimens), 
carbolic  acid  (for  watery  specimens),  and  beechwood  creosote. 
Anilin  oil  will  clear  from  70  or  80  per  cent  alcohol.  It  should  be 
followed  by  oil  of  bergamot,  cassia,  or  wintergreen,  according  to 
McClung.  Clove  oil  should  not  be  used  for  celloidin  sections  because 
it  dissolves  celloidin.  It  is  also  inapplicable  ordinarily  after  most 
anilin  dyes  because  of  its  tendency  to  extract  them.  Among  the 
best  reagents  for  celloidin  sections  are  cedar-wood  oil,  oil  of  origanum, 
creosote,  and  Eycleshymer's  clearer  (memorandum  4,  p.  63). 

While  " clearing"  refers  especially  to  the  rendering  transparent 
of  tissue  elements,  and  dealcoholization  to  the  removal  of  alcohol 
previous  to  imbedding  in  paraffin,  very  frequently  the  same  reagent 
is  used  for  either  purpose  and  the  term  " clearing"  has  come  to  be 

used  in  either  sense. 

MOUNTING 

After  tissues  have  been  cleared  the  final  step  is  to  mount  them 
in  some  suitable  medium  for  preservation  and  inspection. 

If  tissues  are  to  be  mounted  directly  from  water  or  aqueous 
media,  glycerin,  glycerin-jelly,  or  Farrant's  solution  is  used  ordi- 
narily. If  the  alcoholic  dehydration  method  is  employed,  balsam, 
gum  damar,  or  euparal  is  the  final  mounting-medium.  The  balsam 
or  damar  is  dissolved  commonly  in  xylol,  although  turpentine,  chlo- 
roform, or  benzol  may  be  used  as  the  solvent.  In  my  experience 
xylol-balsam  is  the  most  satisfactory  for  ordinary  purposes.  How- 
ever, some  of  our  best  American  technicians  prefer  gum  damar 

dissolved  in  xylol. 

IMBEDDING 

In  order  to  section  tissues  or  objects  satisfactorily  it  is  frequently 
necessary  to  imbed  them  in  a  suitable  matrix.  Simple  imbedding 
consists  in  merely  surrounding  the  object  by  an  appropriate  medium 

f 


General  Statement  of  Methods  23 

to  hold  it  in  place  while  it  is  being  cut.  In  interstitial  imbedding 
the  object  is  saturated  (infiltrated)  with  the  imbedding-substance 
which,  when  all  cavities  and  interstices  are  filled,  is  caused  to  set; 
thus  it  supports  all  parts  of  the  tissue  and  holds  the  components  in 
place  when  sections  are  made.  Infiltration  imbedding  is  of  great 
importance  to  microscopists  and  much  of  the  space  of  the  present 
book  is  given  up  to  drilling  the  student  in  the  details  of  the  two  chief 
infiltration  methods,  viz.,  the  paraffin  method  and  the  celloidin 
method.  Infiltration  with  gum  is  also  not  infrequently  resorted  to, 
especially  for  tissues  which  would  be  injured  by  alcohol,  or  for 
sectioning  by  the  freezing  method. 

Paraffin  is  a  translucent,  waxy  material  derived  from  various 
sources,  one  of  the  commonest  of  which  is  crude  petroleum.  Par- 
affins of  low  and  of  high  melting-points,  termed  respectively  soft  and 
hard  paraffin,  should  be  kept  on  hand  so  that  mixtures  of  different 
degrees  of  hardness  may  be  made  up  as  necessity  demands. 

Celloidin  is  a  form  of  pyroxylin  (guncotton  or  collodion  cotton) 
specially  prepared  for  interstitial  imbedding.  It  is  dissolved  in  a 
mixture  of  ether  and  alcohol  (p.  8,  reagent  4)  and  solutions  of  two 
or  three  strengths  are  used  for  infiltration.  For  details  see  the 
method,  p.  59.  Collodion  instead  of  celloidin  is  used  by  some  workers 
(see  memorandum  11,  p.  65). 

AFFIXING  SECTIONS 

When  mounting  sections  upon  a  slide,  especially  if  they  are 
yet  to  be  stained,  it  is  usually  necessary  to  affix  them  firmly  to  the 
slide  to  prevent  later  displacement.  For  paraffin  sections  Mayer's 
albumen  fixative  (reagent  19,  p.  12),  or  a  combination  of  this  method 
with  the  water  method,  is  most  widely  used.  The  water  method 
alone  often  proves  adequate,  particularly  with  thin  sections.  The 
slide  is  flooded  with  water,  or  better,  albuminized  water  made  by 
adding  3  drops  of  albumen  fixative  to  30  c.c.  of  distilled  water,  and 
the  sections  are  floated  upon  its  surface.  The  paraffin  ribbon  should 
be  gently  heated  until  it  becomes  translucent  but  not  melted,  in  order 
to  make  it  spread  and  flatten  properly.  As  the  layer  of  water 
evaporates,  the  sections  are  slowly  drawn  down  into  close  contact 
with  the  slide.  When  perfectly  dry  they  are  usually  so  firmly  affixed 


24  Animal  Micrology 

that  they  will  not  become  detached  even  after  the  removal  of  paraffin 
from  them.  It  is  common,  however,  and  safer  to  use  a  thin  film  of 
albumen  fixative  as  a  cementing  substance  between  the  water  and 
the  surface  of  the  slide. 

In  the  case  of  celloidiri  sections,  if  only  one  or  a  few  sections  are 
to  be  mounted  on  one  slide,  it  is  a  common  practice  to  stain  the 
sections  and  transfer  them  through  the  various  reagents,  even  to 
clearing,  before  mounting  them  on  the  slide.  In  such  cases  the 
sections  need  not  be  fixed  to  the  slide.  With  serial  sections,  however, 
the  sections  must  be  held  in  place  in  some  way  during  their  transition 
through  the  reagents  (see  memoranda  12  and  13,  p.  165).  Unlike 
paraffin,  the  celloidin  is  not  ordinarily  removed  from  the  tissues. 

DECOLORIZING 

Not  infrequently  in  staining,  the  tissue  becomes  overstained  and 
requires  that  some  of  the  color  be  extracted  from  certain  of  the 
elements  to  bring  about  a  proper  differentiation.  The  fact  that 
certain  tissue  elements  retain  stain  more  tenaciously  than  others  is 
sometimes  taken  advantage  of  and  overstaining  followed  by  decolori- 
zation  is  practiced  intentionally.  Alcohol  slightly  acidulated  with 
hydrochloric  acid  (0.1  to  1  per  cent)  is  commonly  used  for  the 
extraction  of  surplus  color.  In  special  cases  other  decolorizers  are 
used:  for  example,  iron-alum  in  the  iron-hematoxylin  method 
(reagent  17,  p.  10). 

Overstaining  tissue  and  then  partially  decolorizing  it  is  sometimes 
designated  as  regressive  staining  in  contradistinction  to  progressive 
staining  in  which  the  dye,  once  taken  up  by  the  tissue,  is  not  removed. 
In  progressive  staining  differentiation  is  accomplished  through  the 
selective  affinity  of  dyes  for  different  elements. 

BLEACHING 

In  some  cases  tissues  are  obscured  because  of  the  presence  of 
natural  pigments  or  on  account  of  blackening  caused  by  the  fixing 
reagent.  Such  tissues  must  be  bleached.  Chlorine,  peroxide  of 
hydrogen,  or  sulphurous  acid  are  commonly  employed.  A  method 
is  given  in  memorandum  12,  p.  44. 


General  Statement  of  Methods  25 

CORROSION 

To  obtain  skeletal  structures,  as,  for  example,  the  spicules  of 
sponges  or  the  hard  parts  of  insects,  various  methods  of  corrosion 
are  employed.  Nitric  acid,  caustic  potash,  caustic  soda,  eau  de 
Javelle  are  reagents  often  used  for  this  purpose.  Corrosion  prepara- 
tions of  injected  vessels  and  cavities  may  also  be  made. 

DECALCIFICATION  AND  DESILICIDATION 

Tissues  impregnated  with  lime  salts  or  with  silica  must  have 
such  hard  parts  removed  usually  before  they  can  be  sectioned.  For 
decalcification,  one  of  several  acids  may  be  used.  The  details  are 
given  in  the  chapter  on  bone,  tooth,  etc.  (chap.  xi).  For  decalcifying 
reagents,  see  Appendix  B,  v. 

Where  desilicidation  is  necessary  hydrofluoric  acid  may  be 
employed,  although,  because  of  its  property  of  attacking  mucous 
membranes,  its  use  is  attended  with  more  or  less  danger  for  the 
operator.  It  is  added  drop  by  drop  to  the  tissue  which  has  pre- 
viously been  placed  in  a  paraffin-coated  vessel  (the  acid  attacks 
glass).  If  the  tissue  is  not  too  heavily  impregnated  with  silica,  it  is 
safer  to  use  an  old  section  razor  and  try  to  cut  sections  without 
previously  treating  them  with  hydrofluoric  acid. 

INJECTION  METHODS 

The  injection  of  colored  masses  into  the  blood  vessels  and  other 
vessels  of  the  body  is  frequently  practiced  to  aid  in  determining  their 
distribution  and  their  relation  to  the  surrounding  tissues.  The  dye 
is  termed  the  coloring  mass  and  the  substance  to  which  it  is  added  the 
vehicle. 

ISOLATION  OF  fflSTOLOGICAL  ELEMENTS 

Isolation  is  one  of  the  most  valuable  means  of  forming  a  correct 
conception  of  cells  and  fibers.  It  has  the  advantage  over  sections 
that  the  elements  may  be  inspected  in  their  entirety  and  from 
all  sides.  The  separation  is  accomplished,  as  already  noted,  by 
(1)  reagents  which  dissolve  or  soften  cell  cement  and  interstitial 
material  without  seriously  affecting  the  cells  (maceration  or  disso- 
ciation), or  (2)  mechanically  by  means  of  dissecting  needles  (teasing). 


26 


Animal  Micrology 


or  both.  Hardening  and  fixing  reagents  in  general  if  diluted  to 
about  one-tenth  are  efficient  for  dissociation.  Gage  recommends 
normal  saline  as  preferable  to  water  for  dilution.  The  dissecting 
microscope  or  some  kind  of  lens-holder  and  lens  are  valuable  aids 
in  isolating  tissue  elements.  For  practical  methods  consult  chap,  x; 
for  reagents,  Appendix  B,  iv. 

NORMAL  OR  INDIFFERENT  FLUIDS  .FOR  EXAMINING  FRESH 

TISSUES 

It  is  desirable  frequently  to  examine  fresh  material  in  as  near  a 
natural  condition  as  possible,  hence  recourse  is  had  to  the  so-called 
indifferent  fluids.  While  not  wholly  indifferent,  they  ordinarily 
produce  but  slight  changes  in  tissues  and  their  elements  from  the 
viewpoint  of  the  microscopist.  The  liquids  most  commonly  used 
for  this  purpose  are  discussed  in  Appendix  B,  iii. 

GENERAL  SCHEME  FOR  MOUNTING  WHOLE  OBJECTS  (IN  TOTO 
PREPARATIONS)  OR  SECTIONS 


Whole  Objects  (for  balsam  mounts) 
Killing  and  fixing 

Washing 

Staining 

(Decolorizing  if  necessary) 

Dehydrating 

Clearing 


"^Section  Methods  (paraffin  and  celloidin) 
Killing  and  fixing 

Washing 

(Staining,  if  to  be  stained  in  bulk) 
Hardening  and  dehydrating 
Absolute  alcohol 


Mounting 


//  not  stained  in 

bulk 
Through  alcohols 

to  stain 

Staining 
Washing 

Dehydrating  (and 
decolorizing  if 
necessary) 


Paraffin  Method 
Dealcoholization  (xylol) 

Melted  paraffin 
Imbedding 


Sectioning 
Affixing  sections 
Removal  of  paraffin 
Absolute  alcohol 
Clearing 


//  not  stained  in 

bulk 
Staining 

Washing  (and 
decolorizing  if 
necessary) 


Celloidin  Method 
Ether-alcohol 

Thin  celloidin 
Thick  celloidin 
Imbedding 
Sectioning* 

Dehydrating  to  95 
per  cent  alcohol 

Clearing 
Mounting 


Mounting 

*  If  sections  are  to  be  arranged  serially  they  are  best  mounted  in  sheets  or  affixed  to 
the  slide  as  soon  as  cut. 


CHAPTER  III 
KILLING  AND  FIXING 

CAUTIONS. — 1.  Use  only  fresh  tissues  and  work  rapidly  so  that 
the  tissue  elements  will  not  have  time  to  undergo  postmortem  changes. 

2.  Remove  organs  carefully  and  avoid  crushing  or  pressing  the  parts 
to  be  prepared. 

3.  Tissues  should  never  be  allowed  to  dry  from  the  time  they  leave 
the  animal  until  they  are  finally  mounted  for  microscopical  examination 
except  at  one  point  in  the  paraffin  method. 

4.  Use  only  small  'pieces  (2  to  6  mm.  cube)  of  tissue  whenever 
possible,  or  penetration  of  the  reagent  will  be  insufficient.     Embryos 
and  small  objects  up  to  4  cm-  in  size  may  be  placed  entire  in  certain  of 
the  fixing  fluids. 

5.  For  fixing  and  hardening,  the  bulk  of  the  fluid  should  be  from 
10  to  50  times  that  of  the  object.     Too  many  pieces  should  not  be  placed 
in  the  same  'vial. 

6.  Use  only  clean  reagents.    It  is  well  to  let  the  object  rest  on  a  bit 
of  cotton  in  the  bottom  of  the  vial  or  have  it  suspended  from  the  vial 
mouth  so  that  the  reagent  may  penetrate  equally  from  all  sides.    Pene- 
tration is  aided  by  heat. 

7.  When  necessary  to  wash  fresh  tissue,  it  is  usually  best  to  use 
normal  saline,  and  not  water.     Let  it  flow  gently  over  the  surface  of  the 
object  or  slowly  twirl  the  latter  in  the  fluid.     Do  not  scrape  off  foreign 
matter. 

8.  In  many  cases  the  killing  and  fixing  reagent  does  not  harden  the 
tissue  sufficiently  and  the  hardening  process  must  be  completed  in  alcohol. 

9.  Keep  the  reagents  and  preparations  from  direct  sunlight. 

10.  Carefully  label  each  vessel  containing  tissue.    State  the  contents, 
the  fluid  used,  and  the  date.     Label  on  the  side. 

11.  Keep  on  cards  a  careful  record  of  the  reagents  used,  and  the 
time  when  changed,  for  each  separate  piece  of  tissue. 

27 


28  Animal  Micrology 

PRACTICAL  EXERCISE 

Kill  a  frog  or  other  small  vertebrate  by  placing  it  under  a  bell-jar 
which  contains  a  bit  of  cotton  saturated  with  chloroform.  Open  the 
body  as  soon  as  possible  after  death  and  secure  the  tissues  specified 
below. 

1.  Alcohol  Fixation. — Remove  the  dorsal  aorta  and  small  pieces 
of  the  liver  and  harden  in  absolute  alcohol  (at  least,  not  less  than 
95  per  cent)  in  a  vial  or  small  bottle.     The  tissue  will  be  ready  for 
further  treatment  in  two  days. 

Larger  pieces  of  tissue  require  longer  time.  The  pieces  should 
be  thin.  Change  the  alcohol  every  day  for  the  first  three  days. 

Alcohol  is  in  most  instances  an  unsatisfactory  fixing  reagent,  but 
it  is  frequently  employed  because  it  is  usually  at  hand  and  is  easily 
manipulated.  Hot  absolute  alcohol  is  very  often  used  for  insects. 
Acetic  acid  (p.  207)  is  used  with  alcohol  sometimes  to  increase  pene- 
tration and  to  counteract  its  tendency  to  shrink  tissues.  The  mix- 
ture is  usually  preferable  to  alcohol  alone. 

2.  Fixing  with  Zenker's   Fluid. — Place  small  pieces  of  liver, 
kidney,  pancreas,  spleen,  cardiac  and  pyloric  ends  of  the  stomach, 
bladder,  spinal  cord,  and  brain  in  about  ninety  times  their  bulk  of 
Zenker's  fluid.     Remove  a  piece  of  the  intestine  about  12  mm.  long, 
and  after  washing  it  thoroughly  in  normal  saline  place  it  in  a  vial  of 
the  fixing  mixture.     After  fixation,  which  requires  from  6  to  24  hours, 
wash  the  objects  in  running  water  for  from  12  to  24  hours.     Then 
transfer  them  through  35  and  50  per  cent  alcohol  (20  minutes  each) 
into  70  per  cent  alcohol.     Add  sufficient  iodine  solution  to  give  the 
alcohol  a  port-wine  color.     The  iodine  will  remove  any  mercuric  crys- 
tals which  may  have  formed  in  the  tissues.     As  often  as  the  color 
disappears  the  iodine  must  be  renewed.     After  from  12  to  36  hours 
of  this  treatment,  the  color  persists  and  the  objects  should  then  be 
transferred  to  fresh  70  or  80  per  cent  alcohol,  which  must  be  renewed 
until  it  no  longer  extracts  iodine  from  the  specimens.     Too  pro- 
longed washing  with  iodine  solution  tends  to  undo  the  work  of  fixation 
and  to  hinder  staining.     Many  workers  prefer  to  omit  the  treat- 
ment with  iodine  until  the  tissue  has  been  cut  into  sections.     In 
such  cases  slides  bearing  sections  are  treated  with  dilute  iodized 


Killing  and  Fixing  29 

alcohol  for  30  minutes  and  then  washed  thoroughly  in  70  per  cent 
alcohol. 

Zenker's  is  one  of  the  best  general  fixing  agents  known.  Any  of 
a  great  variety  of  stains  may  be  used  after  it,  and  it  fixes  satisfac- 
torily almost  any  kind  of  tissue.  The  time  during  which  objects 
should  be  left  in  the  fluid  varies  from  30  minutes  for  delicate  ones 
to  24  to  36  hours  for  larger  or  denser  tissues,  although  many  objects 
may  be  left  a  longer  time  without  injury. 

3.  Fixing  with  Bouin's  Fluid. — Place  small  pieces  of  trachea, 
tongue,  cornea,  intestine,  and  testis  or  ovary  in  Bouin's  fluid  for 
from  4  to  16  hours.     After  fixation,  wash  the  tissues  in  several  changes 
of  50  per  cent  alcohol,  then  in  70  per  cent  alcohol  until  the  yellow 
color  ceases  to  be  discharged.     Preserve  in  70  or  80  per  cent  alcohol. 
Bouin's  fluid  is  an  excellent  reagent  which  gives  a  very  delicate 
fixation.     It  is  one  of  the  safest  for  the  beginner  because  it  is  almost 
impossible  to  go  wrong  in  its  use.     Objects  may  be  left  a  consider- 
able time  in  it  without  injury. 

4.  Formalin  as  a  Fixing  Reagent. — Place  a  piece  of  spinal  cord, 
sciatic  nerve,  liver,  and  fragments  of  muscle  in  which  nerves  terminate 
in  10  per  cent  formalin  and  leave  until  needed  for  work  lp,ter.     For- 
malin in  varying  percentages  is  widely  used  for  the  preservation  and 
fixation  of  specimens  for  dissection.     It  has  been  employed  especially 
for  the  central  nervous  system.     However,  Miss  Helen  D.  King 
(Journal  of  Comparative  Neurology,  XXIII,  No.  3  [August,  1913]), 
who  has  made  a  careful  study  of  the  effects  of  this  fluid  on  the  brain 
of  the  white  rat,  pronounces  it  unfit  for  cytological  work  because  of 
its  tendency  to  swell  brain  tissue.     On  the  other  hand,  she  finds  that 
nerve  tracts  are  apparently  not  affected  adversely  by  it.     If  a  for- 
malin-fixed brain  is  to  be  used  for  histological  purposes,  she  advises 
its  transfer  to  alcohol  as  soon  as  it  is  fixed  and  hardened.     Formalin 
ordinarily  has  a  slightly  acid  reaction  due  to  the  presence  of  formic 
acid.     Miss  King  finds  such  formalin  less  harmful  than  that  which 
has  been  neutralized.     For  faithful  preservation  of  cells,  however, 
she  prefers  some  other  fixing  fluid,  such  as  Bouin's.     For  simple 
preservation,  solutions  ranging  from  2  to  5  per  cent  are  adequate, 
but  for  fixation   the   solution  should   be   stronger  (10  per  cent). 


30  Animal  Micrology 

Entire  human  brains  may  be  fixed  and  hardened  in  a  10  per  cent 
solution. 

Formalin  is  much  used  for  fixing  and  preserving  when  frozen 
sections  are  to  be  made,  and  it  is  particularly  serviceable  where  a 
study  of  fat  is  desired.  It  also  interferes  less  with  microchemical 
tests  than  most  other  reagents. 

MEMORANDA 

1.  Tissues  Are  Preserved  in  Alcohol  of  from  70  to  85  per  cent  strength, 
but  if  they  are  to  remain  several  months  it  is  better  to  preserve  them  in  a 
mixture  of  equal  parts  of  glycerin,  distilled  water,  and  95  per  cent  alcohol. 

2.  Hardening. — Read  carefully  the  remarks  on  hardening  in  chap.  ii. 

3.  Tissues  Should  Not  Be  Left  in  the  Fixing  Agent  longer,  ordinarily, 
than  is  necessary  to  get  results.     Some,  however,  require  a  long  time  to 
bring  out  the  optical  differences  of  their  elements.     Experience  alone  can 
teach  the  time  required  in  a  given  case.     Such  a  reagent  as  formalin  kills, 
fixes,  hardens,  and  preserves,  all  at  the  same  time.     However,  see  remarks 
under  4,  p.  29. 

4.  Fixation  by  Injection  is  highly  advantageous  with  many  tissues  be- 
cause the  fixing  fluid  is  brought  quickly  into  contact  with  all  parts  of  the  body. 
The  vascular  system  is  first  washed  out  with  normal  salt  solution  and  then 
filled  with  the  fixer.    For  fluids  containing  corrosive  sublimate  a  glass  syringe 
and  cannula,  instead  of  a  metal  one,  should  be  used. 

5.  Hollow  Organs  should  be  filled  with  the  fixing  fluid  and  then  sus- 
pended in  a  vessel  of  the  same. 

6.  For  Transferring  Small  Objects  through  reagents  the  method  of 
Walton  is  an  excellent  one.     For  the  several  reagents  he  uses  shell  vials  which 
measure  about  10  cm.  in  height  by  3  cm.  in  diameter.    Through  the  center 
of  a  flat  cork  which  fits  the  vials  a  hole  is  made  and  a  glass  tube  (about 
9  cm.X  1 . 5  cm.)  is  inserted  so  that  its  lower  end  dips  well  into  the  reagents 
in  the  vials.    The  lower  end  of  the  tube  is  closed  with  fine-meshed  cloth  and 
the  objects  are  placed  within  the  tube.    To  transfer  the  objects  one  simply 
removes  the  cork  bearing  the  tube  and  inserts  it  in  the  vial  containing  the 
desired  reagent.    The  upper  end  of  the  tube  may  be  closed  with  a  cork  of 
the  proper  size.    To  avoid  disturbance  from  changes  in  air  pressure  a  small 
hole  should  be  bored  in  the  side  of  the  tube  just  below  the  lower  level  of  the 
larger  cork.    The  vials  are  supported  as  indicated  in  memorandum  7. 

Very  small  objects  (e.g.,  small  eggs)  may  also  be  made  up  into  little 
packets  in  bits  of  the  cast-off  epidermis  of  the  frog  or  salamander,  according 
to  the  method  of  Professor  Boveri.  The  epidermal  film  is  spread  over  the 
concavity  of  a  hollow  ground  slide  and  saturated  with  alcohol  of  the  same 
strength  as  that  which  surrounds  the  objects.  After  the  latter  have  been 


Killing  and  Fixing  31 

transferred  by  means  of  a  pipette  to  this  sheet  of  epidermal  tissue,  two 
opposite  edges  of  the  sheet  are  folded  over  the  objects,  then  the  other  two 
are  brought  together,  twisted  about  each  other,  and  pinned  with  a  fine  phi. 
The  pin,  bearing  a  label,  is  used  as  a  handle  to  transfer  the  little  bag  of 
objects  from  one  reagent  to  another.  Since  the  epidermal  tissue  cuts  readily, 
it  need  not  be  removed  if  the  objects  are  to  be  imbedded  for  sectioning. 

Professor  C.  E.  McClung  places  fresh  bits  of  tissue  on  small  strips  of 
paper  with  the  proper  index  number  on  the  opposite  side.  The  paper  is 
then  inverted  on  the  surface  of  the  fixing  fluid.  The  tissue  will  adhere  to 
the  paper  through  all  subsequent  processes.  For  washing,  dehydrating, 
etc.,  simply  float  the  paper  on  the  proper  fluid.  A  number  of  objects  may 
thus  be  handled  together.  The  plane  of  section  may  also  be  marked  by 
the  paper. 

Gelatin  capsules,  such  as  are  used  for  medicines,  have  been  recommended 
for  small  objects  by  various  workers.  A  hole,  of  sufficient  size  to  let  through 
the  reagent  but  hold  back  the  objects,  is  pricked  in  each  end  of  the  capsule. 

7.  Shell  Vials,  Small  Bottles,  etc.,  when  in  use  are  best  supported  in 
shallow  auger  holes  of  proper  size  in  thick  blocks  of  wood. 

8.  Material  Which  Is  to  Be  Kept  Indefinitely  should  be  put  in  tightly 
stoppered  vials  in  a  place  away  from  strong  light.    Glass  stoppers  should 
be  used,  since  cork,  besides  shrinking  and  disintegrating  with  age,  may  give 
off  extractives  which  injure  delicate  tissues.    It  is  best  to  pack  the  vials  in  a 
museum  jar  on  cotton  and  then  seal  the  jar  securely  to  prevent  evaporation. 
Material  is  even  more  secure  if  the  museum  jar  is  partly  filled  with  alcohol;  in 
such  a  case  each  small  vial  should  have  a  label  of  the  contents  placed  within  it. 

Another  way  to  prevent  evaporation  from  vials  or  bottles  is  to  "cap" 
them  with  a  suitable  varnish  (see  memorandum  9). 

9.  To  Seal  Bottles  and  Preparation  Jars  ("bottle-capping"),  dip  the 
stopper  and  part  of  the  neck  in  collodion  varnish  made  as  follows: 

Pyroxylin  (e.g.,  collodion  or  celloidin) 1  oz. 

Ether 6  oz. 

Alcohol 8  oz. 

When  the  pyroxylin  has  completely  dissolved  add  2 . 5  drams  of  camphor. 
(From  Pharmaceutical  Era,  XXX,  528.) 

10.  For  the  Preservation  of  Anatomical  Material  for  other  than  cytologi- 
cal  or  histological  dissection,  Professor  George  Wagner  of  our  own  laboratory 
finds  nothing  as  serviceable  as  Keiller's  fluid.    The  formula  is  as  follows: 

Formalin 1.5  parts 

Carbolic  acid 2.5  parts 

Glycerin 10      parts 

Water 86      parts 

If  a  good  grade  of  carbolic  acid  is  used,  disagreeable  odors  will  largely 
be  avoided.  For  embalming,  a  suitable  reservoir  is  filled  with  thp  fluid  and 


32  Animal  Micrology 

suspended  some  3  or  4  feet  above  the  body  to  be  preserved.  About  6  feet 
of  rubber  tubing,  provided  with  clamps  and  a  glass  cannula  of  proper  size 
to  fit  the  femoral  artery,  is  attached  to  the  reservoir.  After  killing  the  animal 
with  illuminating  gas  or  chloroform,  the  fluid  is  injected  through  the  femoral 
artery.  An  important  precaution  is  to  have  the  column  of  fluid  in  the  tube 
and  cannula  free  from  air  bubbles  or  foreign  materials.  The  pressure  should 
be  such  at  all  times  as  to  prevent  blood  from  running  back  through  the 
cannula.  The  animal  should  be  subjected  to  this  embalming  process  for 
two  hours.  Obviously,  by  increasing  the  size  of  the  reservoir  and  the  number 
of  tubes,  several  animals  may  be  treated  at  the  same  time. 

At  the  conclusion  of  the  embalming  process,  if  the  arteries  are  to  be 
injected  with  a  colored  mass,  the  rubber  tube  should  be  disconnected  from 
the  cannula  and  the  latter  be  left  in  place  in  the  artery.  After  the  animal 
has  remained  for  24  hours  in  an  upright  position  the  injection  may  be  under- 
taken. For  study  of  the  blood  vascular  system,  however,  some  workers 
prefer  to  inject  fresh  animals  and  preserve  them  in  formalin.  For  a  satis- 
factory starch  injection  mass  see  memorandum  16,  p.  92. 

In  our  own  laboratory  it  is  the  practice  to  skin  the  embalmed  animal 
and  wrap  the  body  in  cheesecloth  saturated  with  the  embalming  fluid.  It 
is  kept  in  a  zinc  box  of  suitable  size  which  has  a  close-fitting  lid.  We  find 
a  box  25  inches  long,  8  inches  wide,  and  9  inches  deep  a  very  good  size  for 
cats.  For  a  more  detailed  account  of  embalming  with  Keiller's  fluid  see 
Bensley,  Practical  Anatomy  of  the  Rabbit,  pp.  194-98. 

The  following  mixture,  recommended  to  the  author  by  Professor  Kincaid 
of  the  Washington  State  University,  has  given  most  excellent  results.  To  a 
mixture  of  equal  parts  of  glycerin  and  95  per  cent  alcohol  sufficient  formalin 
is  added  to  make  the  whole  about  a  2  per  cent  formalin.  Specimens  remain 
perfectly  flexible  in  this  mixture,  and,  indeed,  after  they  have  become 
thoroughly  saturated,  many  forms  (crustacea,  insects,  etc.)  may  be  removed 
and  kept  as  dry  specimens  which  still  retain  their  flexibility. 

11.  Material  Which  Has  Been  in  Formalin  and  is  to  be  dissected  may 
be  rendered  more  pliable,  and  harmless  to  the  skin,  by  soaking  in  a  3  per  cent 
solution  of  carbolic  acid. 

12.  For  Washing  out  Specimens  it  is  advisable  to  have  the  laboratory 
water-pipe  provided  with  numerous  small  cocks  about  10  cm.  apart,  so  that 
each  student  may  have  the  use  of  one  or  more.    A  piece  of  rubber  tube, 
long  enough  to  reach  to  the  bottom  of  the  vessel  containing  the  specimen  and 
fitted  with  a  bit  of  glass  tube  in  the  free  end,  may  be  attached  to  each  outlet. 
If  the  objects  are  very  small  they  may  be  placed  in  perforated  porcelain 
thimbles  which  may  be  purchased  from  instrument  dealers.    For  minute 
objects  the  thimbles  may  need  to  be  lined  with  fine  gauze.    Bits  of  glass 
tubing  with  fine  gauze  over  the  two  ends  are. also  useful  for  small  objects. 


CHAPTER  IV 

SIMPLE  SECTION  METHODS 
FREE-HAND  SECTION  CUTTING 

This  method  is  important  because  it  requires  no  costly  appli- 
ances; although  the  sections  are  not  as  accurately  cut  as  when 
mechanical  aids  are  used,  the  method  is  simple,  rapid,  and  adequate 
for  the  more  general  histological  and  pathological  work. 

1.  The  section  razor  is  flat  on  one  side  (the  lower)  and  hollow 
ground  on  the  other  (Fig.  15).    It  must  be  sharp. 

2.  A  shallow  glass  dish  or  watch-glass  partly  filled  with  water 
is  also  necessary.    Before  making  a  section,  dip  the  razor  flatwise 
into  the  liquid,  or  use  a  camel's  hair  brush;  see  that  the  upper  surface 
is  well  flooded. 

3.  Sit  in  such  a  way  that  the  forearm  may  be  steadied  against 
the  edge  of  the  table. 

4.  Use  a  piece  of  liver  which  was  fixed  in  formalin,  first  rinsing  it 
in  water.    Take  the  tissue  between  the  thumb  and  forefinger  of  the 
left  hand,  and  hold  it  in  such  a  way  that  a  thin  slice  may  be  cut  by 
drawing  the  knife  along  the  surface  of  the  forefinger. 

5.  Rest  the  flat  surface  of  the  knife  upon  the  forefinger,  and, 
beginning  at  the  heel  of  the  knife,  carefully  draw  the  blade  toward 
you  diagonally  through  the  tissue,  slicing  off  a  thin  section  of  as 
uniform  thickness  as  possible. 

6.  As  each  section  is  cut,  float  it  off  into  the  water;  if  it  adheres 
to  the  blade,  remove  it  by  means  of  a  wet  camel's  hair  brush. 

7.  Practice  until  very  thin  sections  are  obtained,  then  place  the 
dish  upon  a  black  surface,  and  with  a  needle  or  section-lifter  transfer 
the  thinnest  and  best  sections,  if  only  fragments,  to  a  watch-glass 
containing  water. 

NOTE. — In  case  the  tissue  has  b/een  preserved  in  alcohol,  cut  the  sections 
under  70  per  cent  alcohol  instead  of  water,  then  transfer  them  to  50  and  35  per 
cent  alcohol  successively  and  finally  to  water,  leaving  them  in  each  liquid  from 
3  to  5  minutes. 


34  Animal  Micrology 

8.  Next,  place  the  sections  in  about  3  c.c.  of  Delafield's  hema- 
toxylin  diluted  with  an  equal  volume  of  water,  and  leave  them  for 
various  lengths  of  time  (3,  7,  12,  or  more  minutes)  to  determine  the 
time  for  successful  staining. 

9.  Transfer  the  sections  from  the  stain  to  tap  water,  and  gently 
move  them  about  for  from  5  to  10  minutes  to  wash  out  the  excess 
of  the  stain.     If  the  sections  are  still  overstained,  place  them  in 
5  c.c.  of  distilled  water  to  which  3  drops  of  acetic  acid  have  been 
added.     Leave  for  5  minutes,  or  until  they  become  lighter  in  color, 
then  wash  in  several  changes  of  tap  water  until  they  have  again 
become  blue. 

10.  Remove  the  sections  from  the  water  and  transfer  them  through 
35,  50,  70,  83,  and  95  per  cent  alcohol  successively,  leaving  them  from 
3  to  5  minutes  in  each,  and  then  transfer  them  to  absolute  alcohol 
for  10  minutes,  and  finally  to  xylol  for  10  minutes,  or  until  clear. 

11.  Select  one  or  two  of  the  best  sections  and  transfer  them  to 
the  center  of  a  clean  glass  slide.     After  straightening  them  out 
properly,  drain  off  the  excess  of  the  clearer  and  before  the  sec- 
tions can  become  dry,  add  a  drop  of  Canada  balsam.     Carefully 
lower  a  clean  cover-glass  (for  cleaning  see  memorandum  14,  p.  56) 
on  to  the  balsam.    There  should  be  just  sufficient  balsam  to  spread 
evenly  under  the  cover  without  exuding  around  the  edges. 

12.  Label,  stating  card  number,  name  of  the  preparation,  and 
other  data  that  it  is  desired  to  add  (see  p.  48,  step  10). 

13.  Carry  one  of  the  pieces  of  stomach  prepared  in  Zenker  through 
the  same  treatment.     The  sections  should  be  transverse  sections  of 
the  stomach  wall. 

14.  Clean  up  all  dirty  glassware  immediately. 

MEMORANDA 

1.  The  Thinnest  Sections  are  not  always  the  best.    For  a  general  view 
of  an  organ,  large,  comparatively  thick  sections  are  usually  better;  for  details 
of  structure,  thin  sections. 

2.  Small  Pieces  of  Tissue  may  be  cemented  to  a  cork  if  too  small  to  hold 
conveniently  between  thumb  and  forefinger.    A  piece  of  stout  copper  wire 
is  heated  for  a  moment  in  the  flame  and  touched  to  a  bit  of  paraffin.    As  the 
paraffin  melts,  transfer  drops  of  it  to  the  edge  of  the  tissue,  which  has 


Simple  Section  Methods 


35 


been  previously  placed  on  the  cork.  The  paraffin  cools  and  holds  the 
tissue  fast. 

Another  and  better  method  of  handling  a  small  object  is  to  imbed  it  in  a 
piece  of  hardened  liver.  In  sectioning,  the  liver  as  well  as  the  object  is 
sliced,  but  they  readily  separate  when  placed  in  alcohol.  Beef  liver  or  dog 
liver  is  prepared  for  such  purposes  by  hardening  pieces  about  5X2X2  cm. 
in  size  in  95  per  cent  alcohol  for  24  hours,  and  then  transferring  to  fresh 
95  per  cent  alcohol  until  needed.  When  much  hand  sectioning  is  to  be  done, 
a  supply  of  hardened  liver  should  be  kept  on  hand. 
Many  small  objects  may  be  held  between  pieces  of 
pith  and  successfully  sectioned. 

3.  Well  Microtomes  (Fig.  27)  are  inexpensive 
instruments  which  are  used  for  simple  sectioning. 
Such  a  microtome  consists  of  a  tube  in  which  the 
object  is  placed,  and  at  one  end  of  which  is  a  plate 
to  guide  the  razor.    The  other  end  is  provided 
with  a  screw,  which,  when  turned,  pushes  the  con- 
tents of  the  tube  above  the  plate,  thus  making  it 
possible  to  cut  sections  of  a  uniform  thickness. 
The  object  to  be  cut  must  be  firmly  fixed  in  the 
well.    Such  tissues  as  kidney,  liver,  spleen,  hard 
tumors,  cartilage,  etc.,  may  be  held  sufficiently 
rigid  by  wedging  small  slabs  of  carrot,  turnip,  pith, 
or  hardened  liver  in  about  them.    These  support- 
ing  substances   must,   of   course,   rest  squarely 
against  the  bottom  of  the  well.    Soft  tissues,  such 
as  soft  tumors  or  brain,  must  be  imbedded.    Three 
parts  of  paraffin  and  one  part  of  vaseline  melted 
together  and  thoroughly  mixed  makes  a  very  good 

imbedding-mass  for  a  w^ell  microtome.  To  imbed,  warm  the  microtome 
slightly  and  fill  the  well  with  the  imbedding-mixture.  Remove  all  liquid 
from  the  surface  of  the  tissue,  and  pass  it  below  the  surface  of  the  mixture 
just  as  it  begins  to  harden  around  the  edges.  When  the  imbedding-mass 
has  become  cold  the  sections  are  cut  in  the  ordinary  way. 

4.  Temporary  Mounts  may  be  made  directly  from  water  after  staining 
by  using  glycerin  as  a  mounting-medium.    Transfer  the  section  to  the  slide, 
add  a  drop  or  two  of  glycerin,  and  a  clean  cover-glass. 


FIG.  27.— Well  Microtome 


CHAPTER  V 

THE  PARAFFIN  METHOD:  INFILTRATION  AND 
SECTIONING 

1.  From  70  per  cent  alcohol  take  a  small  piece  of  intestine 
(6  mm.  long)  fixed  in  Zenker's  fluid,  and  also  pieces  of  other  tissues 
fixed  in  this  fluid,  and  proceed  according  to  the  following  schedule. 
Keep  accurate  records  on  your  cards. 

2.  Immerse  in  95  per  cent  alcohol  for  30  to  45  minutes.     A  longer 
time  will  do  no  harm. 

3.  Transfer  to  absolute  alcohol,  1  hour.     Before  transferring  to 
absolute,  remove  the  excess  of  95  per  cent  alcohol  from  the  object 
by  touching  it  with  a  piece  of  blotting  paper  or  a  clean  cloth,  but  do 
not  let-- it  become  dry. 

4.  Xylol  and  absolute  alcohol  equal  parts,  30  minutes. 

5.  Xylol,   1J  hours,  or  until  the  object  looks  clear.     Rapidly 
remove  all  excess  of  xylol  before  proceeding  with  step  5,  but  do  not 
allow  the  tissue  to  become  dry  or  dull  looking.     (For  delicate  objects 
see  p.  53.) 

6.  Melted  paraffin  (melting-point  about  50°  C.),  2  hours.     The 
object  may  be  left  an  hour  longer,  but  it  is  best  to  avoid  as  much  as 
possible  subjecting  tissues  to  an  elevated  temperature.     Shift  its 
position  in  the  paraffin  once  or  twice  to  facilitate  penetration  of  the 
latter. 

The  duration  of  the  paraffin  bath  varies  according  to  the  size  and 
density  of  the  object.  Many  objects  of  from  3  to  5  mm.  in  thickness 
are  thoroughly  saturated  within  an  hour  or  less;  others  which  are 
more  impervious  or  which  have  impenetrable  coverings  may  require 
several  hours  or  even  days.  Lee,  assuming  that  melted  paraffin 
will  penetrate  as  quickly  as  cold  oil,  takes  as  a  guide  the  length  of 
time  required  to  clear  the  object  in  cedar  oil. 

CAUTIONS. — a)  Do  not  have  the  bath  too  hot.  Cooked  tissues  are 
worse  than  useless. 

36 


The  Paraffin  Method  37 

b)  To  keep  material  clean,  it  is  well  to  have  a  false  bottom  of 
paper  in  the  vessel  containing  paraffin.  Make  this  by  swinging  a 
strip  of  white  paper  into  the  cup  so  that  the  loop  of  the  paper  is 
submerged  in  paraffin  and  the  ends  attached  on  either  side  to  the 
mouth  of  the  cup. 

7.  Prepare  paper  boxes  according  to  the  following  instructions: 

A  small  rectangular  block  of  wood  or  a  stick  with  a  flat  end  measuring 
approximately  15X20  mm.  is  used.  Cut  a  strip  of  stiff  glazed  paper  so  that 
it  measures  about  4X7  cm.  Place  the  flat  end  of  the  block  in  the  center  of 
the  paper  with  its  long  diameter  coinciding  with  the  long  diameter  of  the 
paper.  Fold  the  narrow  side  margins  of  the  paper  up  along  the  sides  of 
the  block  first,  then  do  likewise  with  the  ends  of  the  paper.  Turn  the  ears 
which  have  been  formed  at  each  corner  back  over  what  is  to  be  the  end 
of  the  box,  and  then  fold  the  long  end  of  the  paper  back  to  hold  the 
ears  in  place,  and  also  to  make  the  end  of  the  box  of  the  same  height  as 
the  sides.  Manifestly,  any  size  of  box  may  be  made  by  varying  the  size 
of  the  block.  With  a  little  practice  the  same  kind  of  box  may  be  folded 
without  the  use  of  a  wooden  block.  See,  however,  memorandum  14,  p.  45. 

8.  With  a  warm,  wide-mouthed  pipette  transfer  sufficient  melted 
paraffin  to  a  paper  box  to  cover  the  bottom,  then,  with  warm  forceps, 
remove  the  tissue  to  the  box.     Next,  fill  the  box  with  melted  paraffin. 
Orient  the  object  with  heated  needles  if  necessary.     As  soon  as  the 
paraffin  has  congealed  sufficiently  for  the  surface  to  become  opaque, 
cool  it  rapidly  by  plunging  it  into  cold  water;  otherwise  the  paraffin 
will  crystallize  and  become  unsuited  for  sectioning.     Many  workers 
prefer  alcohol  instead  of  water  for  hardening  the  paraffin  block. 
Waste  alcohol  may  be  saved  for  this  purpose. 

CAUTIONS. — a)  Tissues  must  be  oriented  (i.e.,  placed  in  proper 
position  for  cutting)  while  the  paraffin  is  still  in  liquid  condition. 
Arrange  the  tissue  so  that  it  will  be  cut  at  right  angles  (transverse)  or 
parallel  to  the  surface  of  the  organ.  Avoid  oblique  sections  as  they 
are  very  puzzling.  For  present  purposes  of  practice  cut  transverse 
sections. 

6)  If  whitish-looking  patches  are  present  in  the  block  after 
imbedding,  they  are  probably  due  to  xylol  which  has  been  carried 
over  into  the  paraffin.  If  they  occur  in  the  immediate  vicinity  of  the 
object,  the  block  should  be  placed  in  the  bath  again  until  melted,  and 
the  object  should  be  reimbedded. 


38 


Animal  Micrology 


c)  Be  sure  that  every  piece  of  tissue  is  marked  after  it  is  imbedded. 
Tissues  are  sometimes  kept  in  paraffin  for  months  or  even  years 
before  they  are  finally  sectioned.  To  mark,  scratch  the  number  of 
the  record  card  in  the  paraffin  or,  better,  write  it  on  the  paper  box 
and  leave  the  box  in  place. 

CUTTING  SECTIONS 

9.  Study  the  paraffin  microtome  (e.g.,  Fig.  28  or  Fig.  29) ;  identify 
the  parts  and  learn  how  the  thickness  of  sections  is  controlled. 


PIG.  28. — Minot  Automatic  Rotary  Microtome 

The  object-carrier  is  adjustable  in  three  planes  and  is  perfectly  rigid.  The  knife- 
carrier  is  also  adjustable  and  extra  heavy  and  solid.  The  feed  is  controlled  by  an  adjust- 
able cam,  giving  cuts  of  any  number  of  microns  in  thickness  from  1  to  25.  By  means  of 
an  automatically  closing  split-nut  the  carriage  is  returned  to  the  beginning  position  after 
the  screw  is  fed  out  the  entire  length.  For  details  of  construction  see  catalogue  of  the 
Bausch  &  Lomb  Optical  Co. 

10.  Proceed  with  the  block  of  paraffin  containing  the  intestine. 
Make  it  fast  to  the  carrying  disk  of  the  microtome  in  the  following 
manner:  Remove  the  disk  from  the  machine  and,  by  means  of  a 


The  Paraffin  Method  39 

heated  steel  spatula  or  copper  wire  flattened  at  one  end,  melt  a 
small  chip  of  paraffin  on  to  it.  Likewise  warm  the  end  of  the 
paraffin  block  and  quickly  press  it  into  the  melted  paraffin  on  the 
disk.  Cement  it  firmly  in  place  by  means  of  the  heated  wire  or 
spatula  and  cool  in  water. 

11.  With  a  sharp  scalpel  trim  the  free  end  of  the  block  so  that 
it  presents  a  perfectly  rectangular  outline  (however,  see  caution  c). 
The  length  should  exceed  the  breadth  by  at  least  one-fourth. 


FIG.  29. — Spencer  Rotary  Microtome 

By  means  of  a  wheel  at  the  back  the  feed  may  be  set  for  any  thickness  from  1  to 
60  microns. 

CAUTIONS. — a)  In  trimming  do  not  cut  farther  back  than  the 
base  of  the  object.  This  leaves  a  wide  shoulder  for  support. 

6)  Leave  a  margin  of  about  2  mm.  around  the  object. 

c)  To  avoid  reversing  sections  in  mounting,  it  is  frequently 
advantageous  to  have  the  imbedding-mass  trimmed  unsymmetri- 
cally.  The  edge  which  first  comes  in  contact  with  the  knife  is  left 
longer  than  the  opposite  edge.  One  may  thus  readily  discover  when 
a  section  or  part  of  a  series  has  been  turned  over. 

12.  Mount  the  object  firmly  in  the  microtome.  It  should  just 
clear  the  knife.  The  flat  end-surface  of  the  paraffin  block  should 


40  Animal  Micrology 

be  parallel  to  the  edge  of  the  knife,  and  the  block  so  oriented  that,  in 
cutting,  the  long  edge  will  meet  the  edge  of  the  knife  squarely. 

13.  Place  the  knife  in  position  with  the  handle  to  the  side  away 
from  the  wheel  (if  a  rotary  microtome  is  used).     By  means  of  the 
adjusting  screws  tilt  the  cutting  edge  slightly  toward  the  object  so 
that  the  side  of  the  knife  will  not  remain  in  contact  with  the  paraffin 
block  after  a  section  has  been  cut.     If  the  knife  has  a  flat  under  sur- 
face it  requires  more  tilt  than  if  the  surface  is  hollow  ground.     For  a 
flat  under  surface  the  tilt  should  be  about  9  degrees  from  the  per- 
pendicular.    See  that  the  knife  is  held  firmly  in  place. 

CAUTION. — The  knife  should  be  kept  in  its  case  when  not  in 
the  machine.  The  edge  is  very  easily  injured. 

14.  Set  the  regulator  so  that  the  microtome  will  cut  sections 
about  10  microns  thick.     A  micron  is  one-thousandth  of  a  milli- 
meter. 

15.  Unloose  the  catch  which  locks  the  wheel  and  revolve  the 
wheel  with  the  right  hand.     A  few  revolutions  should  bring  the 
block  of  paraffin  into  contact  with  the  knife.     As  each  new  section 
is  cut,  it  displaces  the  last  one  and  if  the  paraffin  is  of  the  proper 
consistency  unites  by  one  edge  with  the  displaced  section.     Thus  a 
ribbon  or  chain  is  formed.     When  the  ribbon  becomes  of  sufficient 
length,  support  the  free  end  by  means  of  a  hair  brush  held  in  the  left 
hand.     To  prevent  breaking  the  ribbon  avoid  pulling  it  taut.     Vari- 
ous ribbon-carriers  have  been  devised  for  attachment  to  the  micro- 
tome.    The  best  of  these  is  the  cylindrical  carrier  made  by  the 
Spencer  Lens  Co.  according  to  the  suggestions  of  Dr.  C.  E.  McClung. 
For  an  inexpensive,  home-made  form  of  this  see  Hance,  Anatomical 
Record,  X,  No.  8  (June,  1916). 

CAUTION. — Never  bring  a  needle  or  other  hard  object  near  the 
edge  of  the  knife.  If  the  paraffin  does  not  ribbon  properly,  consult 
the  table  at  the  end  of  this  chapter. 

16.  When  a  sufficient  number  of  sections  have  been  cut,  care- 
fully place  the  ribbon  on  a  piece  of  paper.     Protect  it  from  draughts 
of  air  which  will  carry  away  or  disarrange  the  sections. 

17.  Cut  the  ribbon  into  strips  of  such  length  that  they  may  be 
placed  in  successive  rows  one  above  the  other  under  the  cover-glass 


The  Paraffin  Method  41 

that  is  to  be  used.  Allow  one-fourth  for  the  expansion  of  the  ribbon 
when  heated.  Mark  out  on  a  sheet  of  paper  the  exact  size  of  the 
cover-glass  so  that  there  can  be  no  mistake  in  cutting  strips  of 
the  proper  length.  A  margin  of  2  or  3  mm.  should  be  allowed  for 
the  cover. 

18.  Place  a  small  drop  of  albumen  fixative  on  a  clean  glass  slide 
(for  cleaning  see  memorandum  14,  p.  156),  and  spread  it  evenly  over 
the  surface,  except  the  end  which  is  to  bear  the  label  (see  step  10, 
p.  49).    With  a  clean  finger  rub  off  all  of  the  fixative  that  can  be 
easily  removed  so  that  only  a  very  thin  film  remains. 

19.  Flood  the  slide  with  a  few  drops  of  distilled  or  albuminized 
(p.  23)  water  until  the  entire  surface  bearing  the  fixative  is  covered 
by  a  thin  layer  of  water,  but  do  not  put  on  sufficient  to  overflow 
the  edge.     Some  workers  use  albuminized  water  alone  for  affixing 
sections. 

20.  Take  up  the  first  strip  of  paraffin  ribbon  with  a  brush  or 
needle  and  float  it  on  to  the  surface  of  the  water.     The  first  section 
of  the  series  should  be  in  the  upper  left-hand  corner,  but  back  at 
least  10  mm.  from  the  end  of  the  slide.     In  case  the  label  is  to  be 
placed  on  the  left  end  of  the  slide,  allowance  must  be  made  for  it, 
of  course.     Add  the  successive  strips  of  the  ribbon  in  the  order  of 
the  lines  of  a  printed  page  until  as  many  rows  are  in  place  as  will 
conveniently  lie  under  the  cover  (see  step  17,  p.  40),  allowing  for  the 
proper  margins.     See  that  each  section  presents  the  same  aspect 
to  the  observer  as  its  predecessor  (see  step  11,  c,  p.  39). 

21.  Warm  the  slide  gently  until  the  paraffin  flattens  out  and 
becomes  free  from  wrinkles.     Be  careful  not  to  melt  the  paraffin, 
for  heat  sufficient  to  do  so  will  render  the  albumen  useless.     It  is 
safer  to  heat  the  slide  by  placing  it  upon  the  warm  paraffin  oven  for 
a  few  minutes,  instead  of  holding  it  above  a  flame. 

22.  Drain  off  the  excess  of  water  and  set  the  slide  away  to  dry 
after  properly  numbering  it  with  your  glass-marking  pencil.     As 
the  water  evaporates,  the  sections  are  drawn  down  tightly  into  the 
film  of  fixative.     If,  after  drying,  air  is  present  under  the  sections 
it  may  be  seen  from  the  glass  side  of  the  slide.     The  slide  is  seldom 
sufficiently  dried  under  6  hours.     It  is  well  to  leave  it  12  hours;  it 


42  Animal  Micrology 

may  be  left  indefinitely.  The  time  may  be  shortened  by  placing  a 
few  thicknesses  of  blotting  paper  under  the  slide  and  drying  it  on  the 
paraffin  oven  or  in  an  incubator.  Unless  the  slide  is  perfectly  dry 
and  the  ribbon  fully  spread,  the  sections  will  float  off  during  sub- 
sequent treatment.  Take  precautions  to  prevent  particles  of  dirt 
from  settling  upon  the  surface  of  the  sections.  This  is  usually  accom- 
plished by  placing  the  slides  upon  some  kind  of  a  rack  and  covering 
them  with  a  bell-jar.  Prepare  several  other  slides  in  the  same  manner 
as  the  above  if  sufficient  of  the  ribbon  remains. 

NOTE. — As  time  permits,  cut  the  other  sections  which  are  imbedded  in 
paraffin.  When,  as  in  the  present  case,  it  is  not  necessary  to  have  a  complete 
series  of  sections,  you  may  place  fewer  sections  on  a  slide  and  use  smaller  covers. 

When  a  small  cover  is  to  be  used,  place  the  sections  at  the  center  of  the 
slide.  The  center  may  readily  be  determined  by  drawing  the  outline  of  a  slide 
on  a  card  and  connecting  the  opposite  corners  of  the  figure  by  means  of  diagonal 
lines.  When  mounting,  place  a  slide  over  the  diagram;  the  intersection  of  the 
diagonals  shows  the  center. 

At  this  point  the  student  should  make  a  careful  study  of  Appendix  A 
if  he  is  not  already  thoroughly  acquainted  with  the  optical  principles 
involved  in  microscopy. 

MEMORANDA 

1.  If  Paraffin  Becomes  Dirty  it  should  be  melted  and  filtered  through  a 
heated  metal  funnel. 

2.  Oil  of  Cedar,  if  used  for  dealcoholization  before  imbedding,  should 
be  followed  by  at  least  two  changes  of  paraffin  or  the  paraffin  does  not 
thoroughly  replace  the  oil  and  the  object  is  likely  to  drop  out  of  the  sections 
as  they  are  cut.    In  my  experience  this  is  the  commonest  difficulty  which 
beginners  encounter  if  they  use  cedar  oil  for  dealcoholization.    For  this 
reason  xylol  or  chloroform  is  recommended  as  preferable  for  general  work. 

3.  Objects  Imbedded  in  Paraffin  may  be  preserved  in  that  form  indefi- 
nitely.   It  is  one  of  the  most  convenient  ways,  in  fact,  of  preserving  material 
which  is  to  be  sectioned  in  paraffin. 

4.  Small  White  Objects,  if  not  stained  before  imbedding,  should  be 
tinged  with  a  dilute  solution  of  Congo  red  to  facilitate  orientation.    For 
orientation  in  general  see  p.  126,  memorandum  1. 

5.  With  Delicate  Tissues  it  is  necessary  that  the  transition  from  alco- 
hol to  clearer  be  gradual,  hence  it  is  best  to  add  the  clearer,  a  little  at  a 
time,  to  the  last  alcohol,  transferring  it  with'  a  pipette  to  the  bottom  of  the 
alcohol.     See  also  "drop"  method,  pp.  18  and  152. 


The  Paraffin  Method  43 

6.  The  Temperature  of  the  Laboratory  must  be  taken  into  account  when 
sectioning  in  paraffin.     In  summer  use  a  harder,  in  winter  a  softer,  paraffin. 

7.  For  Thin  Sections  use  a  hard  paraffin,  for  thick  sections,  a  softer 
paraffin. 

8.  For  Valuable  Tissues  Which  Crumble  in  Paraffin  Alone  the  following 
somewhat  tedious  process  (Mark,  American  Naturalist  [1885],  p.  628)  may 
be  resorted  to.    Prepare  a  very  fluid  collodion  in  ether-alcohol  and  coat  the 
exposed  surface  of  the  object  immediately  before  cutting  each  section.    If 
the  collodion  leaves  a  shiny  surface  or  produces  a  membrane  when  applied 
to  the  paraffin,  it  is  not  thin  enough  and  must  be  further  diluted  with  ether- 
alcohol.    Apply  the  collodion  with  a  brush  with  all  excess  of  the  fluid  wiped 
away  so  that  the  brush  is  just  moist.    The  fluid  should  touch  only  the  face 
of  the  block  in  which  the  object  is  exposed.    After  applying,  wait  a  few  sec- 
onds for  the  solution  to  dry  before  cutting.    See  also  memorandum  9. 

9.  Johnson's  Paraffin-Asphalt-Rubber  Method  for  brittle  objects  is  a 
very  useful  one.    One  part  of  crude  India  rubber  cut  into  very  small  pieces 
is  mixed  with  99  parts  of  hard  paraffin  which  has  previously  been  melted  and 
tinged  to  a  light  amber  color  with  a  small  amount  of  asphalt  ("mineral 
rubber").    The  mixture  is  then  subjected  to  a  temperature  of  100°  C.  (not 
higher)  for  24  to  48  hours,  or  left  in  a  paraffin  oven  at  60°  C.  for  several  days. 
Use  only  the  supernatant  fluid.    It  is  allowed  to  cool  and  remain  cold  until 
needed,  because  the  rubber  separates  out  after  a  time  if  the  mixture  continues 
melted.    Johnson  (Journal  of  Applied  Microscopy,  VI,  2662)  recommends 
it  as  even  better  than  paraffin  for  all  kinds  of  work  for  which  paraffin  is 
commonly  employed.    Proceed  as  in  the  ordinary  method,  using  xylol  (not 
cedar  oil)  for  dealcoholization  and  also  for  clearing  sections. 

10.  Keep  All  Parts  of  the  Microtome  clean  and  well  oiled  with  watch 
oil  or  pure  paraffin  oil  of  25°  C.    The  instrument  should  be  covered  when  not 
in  use. 

1 1 .  Keep  the  Microtome  Knife  Sharp.    It  should  receive  frequent  strop- 
pings.     For  sharpening  the  knife  two  hones  are  commonly  used. 

Honing. — If  the  knife  is  very  dull  it  is  first  honed  on  a  Belgian  yellow 
hone,  an  open-grained  stone  which  cuts  the  metal  of  the  knife  rapidly. 
The  surface  of  the  stone  is  kept  moist  with  filtered  kerosene  oil  or  lathered 
with  palm-oil  soap.  After  the  nicks  and  other  inequalities  of  the  edge  of 
the  knife  have  been  removed,  the  honing  is  best  finished  on  a  good  fine- 
grained blue-water  stone. 

In  honing,  the  stone  is  laid  flat  on  the  table  with  its  end  toward  the 
operator  and  its  surface  properly  lubricated.  A  very  dull  knife  is  ground 
at  first  on  the  concave  side  only  until  it  develops  a  fine  "wire  edge"  along 
the  full  length  of  the  blade.  It  is  then  ground  on  each  side  alternately 
until  the  wire  edge  has  disappeared  completely.  In  grinding,  the  knife 
must  remain  flat  on  the  hone  and  pass  lightly  over  the  full  length  of  the 


44  Animal  Micrology 

surface,  edge  foremost  in  a  diagonal  direction  from  point  to  heel,  although 
itself  remaining  at  right  angles  to  the  long  axis  of  the  hone.  The  honing 
has  been  sufficient  when  all  nicks  and  wire  edges  have  disappeared  and  the 
knife,  instead  of  catching  and  hanging  when  the  edge  is  drawn  lightly  across 
the  ball  of  the  thumb,  freely  enters  the  moist  epidermis.  Finally  the  blade 
is  wiped  clean  with  a  soft  cloth,  great  care  being  taken  not  to  injure  the  edge. 

Some  workers  prefer  to  use  first  a  yellow  Belgian  hone  wetted  with  a 
moderately  thick  soap  solution,  then  an  Arkansas  stone  with  a  thin  oil. 
Hardesty  (Laboratory  Guide  for  Histology,  p.  186)  uses  only  a  white  Arkansas 
stone  with  oil. 

Stropping. — A  broad  firm  strop  of  finest  calfskin  is  best.  It  should  be 
affixed  to  a  solid  back  so  that  it  will  not  spring  and  thus  round  off  the  delicate 
edge  of  the  knife. 

In  stropping,  the  motions  are  the  same  as  in  honing  (both  sides  of  blade), 
only  the  knife  passes  back  foremost  and  from  heel  to  point.  The  blade 
must  move  lightly  over  the  surface  of  the  strop  with  very  slight  pressure 
on  the  part  of  the  operator.  The  stropping  is  ordinarily  considered  sufficient 
when  the  blade  will  cut  a  loose  hair  freely  along  every  part  of  the  edge. 
An  examination  under  a  low  power  of  the  microscope  should  reveal  no  nicks 
in  the  edge. 

12.  To  Remove  Pigments  and  to  Bleach  Osmic  and  Chromic  Acid 
Materials,  a  3  per  cent  solution  of  peroxide  of  hydrogen  frequently  is  suffi- 
cient.   Tissues  left  too  long  in  this  liquid 
macerate.. 

Mayer's  chlorine  method  is  one  of  the 
best  for  bleaching.     To  several  crystals  of 
^  t;I  chlorate  of  potash  in  a  glass  tube  a  few 

V  \        drops    of    hydrochloric    acid   is   added. 

V  ;>-       %,     When    the    greenish   fumes   of   chlorine 

^     appear,  add  from  5  to  10  c.c.  of  50  per 
FIG.  so.-Metai  L'S  for  molding    cent  alcohol.    The  object,  which  in  the 

imbedding-masses.  7 

meantime  has  been  standing  in  70  per 

cent  alcohol,  is  transferred  to  the  tube.  From  15  minutes  to  24  hours  are 
required  for  bleaching,  depending  upon  the  nature  of  the  material.  It  is 
well  to  suspend  the  object  from  the  mouth  of  the  bottle.  Sections  on  the 
slide  may  be  bleached  in  a  few  minutes.  This  method  is  especially  recom- 
mended for  removing  natural  pigments  and  for  bleaching  osmic  material. 

13.  Large  Objects  May  Be  Cut  in  Paraffin  better  with  a  slanting  knife 
than  with  a  square-set  one.    The  block  of  paraffin  must  be  trimmed  to  a 
three-sided  prism  with  its  most  acute  angle  farthest  from  the  object.    A 
sliding  microtome  is  used  ordinarily  and  the  block  of  paraffin  is  so  oriented 
that  the  knife  enters  at  the  sharpest  angle  of  the  prism.    Each  section  as 
cut  is  removed  with  a  brush. 


The  Paraffin  Method  45 

14.  Metal  or  Porcelain  "L's"  Are  Frequently  Used  Instead  of  Paper 
Boxes  for  molding  paraffin  blocks.  The  two  L's  (Fig.  30)  may  be  placed 
together  on  a  small  glass  or  metal  plate  in  such  a  way  as  to  mold  blocks  of 
any  desired  size.  Before  pouring  the  melted  paraffin  in,  the  inner  walls  of 
the  metal  pieces  should  be  lightly  smeared  with  glycerin  so  that  the  block 
of  paraffin  will  easily  separate  from  them  when  cool.  Many  workers, 
particularly  for  small  objects,  imbed  in  a  watch-glass  and  harden  under 
alcohol. 

DIFFICULTIES  LIKELY  TO  BE  ENCOUNTERED  IN  SECTIONING  IN 
PARAFFIN,  AND  THE  PROBABLE  REMEDY 

1.  Crooked  Ribbons. — a)  Usually  caused  by  wedge-shaped  sections. 
Correct  by  trimming  the  block  of  paraffin  so  that  the  edge  which  strikes 
the  knife  first  and  the  edge  on  the  opposite  side  are  strictly  parallel.    See 
that  the  block  strikes  the  knife  exactly  at  right  angles. 

b)  The  paraffin  may  be  softer  at  one  end  of  the  block  than  at  the  other. 
This  can  be  corrected  only  by  imbedding  the  object  over  again  in  a  homo- 
geneous paraffin. 

2.  The  Object  Makes  a  Scratching  Noise  on  the  Knife  or  Cuts  with  a 
Gritty  Feeling  and  the  sections  perhaps  crumble  and  tear  out  from  the  paraffin. 

a)  This  is  generally  caused  by  too  high  heating  of  the  object  while  in 
the  paraffin  oven.  Not  only  is  such  an  object  worthless,  but  it  endangers 
the  edge  of  the  microtome  knife.  Correct  by  limiting  the  bath  in  paraffin 
to  the  minimum  time  necessary  for  a  proper  penetration  of  the  object  and 
keeping  the  temperature  barely  above  the  melting-point  of  the  paraffin. 

6)  The  fixing  reagent  has  formed  crystals  (e.g.,  corrosive  sublimate) 
which  have  not  been  thoroughly  washed  out. 

See  also  5  (p.  46). 

3.  The  Sections  Wrinkle  or  Jam  Together;  the  object  itself  may  be  com- 
pressed before  the  knife.    This  is  a  serious  fault  because  the  arrangement 
of  the  parts  of  a  tissue  is  greatly  deranged.     It  may  be  due  to  various  causes. 

a)  The  microtome  knife  may  be  dull.  Examine  the  knife  and  sharpen 
it  if_ necessary. 

6)  The  paraffin  may  be  too  soft.  To  remedy  this  defect  employ  one 
or  more  of  the  following  means:  (1)  cool  the  paraffin  block  in  water;  (2)  cut 
the  sections  in  a  cooler  room;  (3)  cut  the  sections  thicker;  (4)  reimbed  in 
harder  paraffin.  See  also  memorandum  11,  p.  47.  If  sections  are  not  too 
badly  wrinkled  they  may  be  flattened  out  by  warming  on  water  as  directed 
in  steps  19-21,  p.  41. 

c)  A  possible  reason  is  that  the  tilt  of  the  knife  is  insufficient  (see  step 
13,  p.  40). 

d)  The  edge  of  the  knife  may  be  smeared  with  a  layer  of  paraffin.     Clean 
the  edge  with  a  cloth  moistened  in  xylol. 


46    *  Animal  Micrology 

4.  The  Sections  Roll  and  Refuse  to  Ribbon. — This  is  one  of  the  most 
exasperating  of  all  defects.    If  the  sections  are  not  tightly  curled,  they 
frequently  unroll  when  placed  on  warm  water  (step  19,  p.  41).    Various 
mechanical  devices  have  been  constructed  to  prevent  this  evil,  but  most 
of  them  are  impractical.    Sometimes  when  a  section  begins  to  roll,  if  the 
edge  is  held  down  by  means  of  a  flat-pointed  hair  brush  the  curling  can  be 
overcome.    If  a  ribbon  can  once  be  started,  the  difficulty  is  frequently 
corrected.    The  sections  should  be  cut  rapidly. 

a)  The  commonest  cause  of  rolling  is  the  hardness  of  the  paraffin. 
This   may   sometimes   be   remedied   by   one   or   more   of   the   following 
means:     (1)    warming    the    knife    with    the   breath;     (2)    cutting   in   a 
warmer  room;    (3)  placing  a  lamp  or  burner  near  the  imbedded  object; 
(4)   warming  the   knife  very  carefully  by  holding  the  back  on  a  warm 
paraffin  bath;  (5)  cutting  the  sections  thinner;   (6)  reimbedding  the  object 
in  softer  paraffin;    (7)   dipping  the  end  of  the  block  in  melted  softer 
paraffin. 

b)  The  tilt  of  the  knife  may  be  too  great  (step  13,  p.  40). 

c)  The  knife  may  be  dull. 

5.  The    Sections   Split   Longitudinally   or   Are    Crossed   by   Parallel 
Scratches. — a)  Look  for  a  nick  in  the  edge  of  the  knife.     Cut  in  a  new  place 
on  the  knife  or  sharpen  it. 

6)  A  bit  of  grit  may  have  gotten  into  the  object  or  the  paraffin,  or  there 
may  be  a  hole  in  the  paraffin.  Reimbed  after  carefully  cleaning  the  object 
in  the  clearing  fluid. 

c)  Tissues  may  contain  hard  substances   (lime  salts,  silica,  crystals 
precipitated  from  fixing  reagents)  which  have  been  imperfectly  washed 
out.    It  is  best  to  take  an  entirely  new  piece  of  tissue  in  which  these  defects 
do  not  exist. 

d)  The  tilt  of  the  knife  may  be  too  great  (step  13,  p.  40),  or  the  trouble 
may  be  due  to  loose  paraffin  on  the  edge  of  the  knife. 

e)  The  object  may  be  too  large  to  cut  in  paraffin.    Try  smaller  pieces 
of  tissue  or  use  the  celloidin  method. 

6.  The  Knife  Scrapes  or  Rings  as  It  Passes  Back  over  the  object  after 
having  cut  a  section. 

a)  This  is  sometimes  caused  by  a  knife  with  either  too  great  or  too  little 
tilt  (step  13,  p.  40). 

6)  The  object  may  be  too  tough  or  hard  to  cut  in  paraffin  without 
springing  the  edge  of  the  knife  (see  7,  6,  p.  47). 

c)  The  blade  of  the  knife  may  be  too  thin. 

7.  The  Sections  Vary  in  Thickness;  the  machine  cuts  one  thick  and  one 
thin  or  misses  a  section. 

a)  This  may  be  caused  by  the  imperfect  mechanical  construction  of  the 
machine.  Old  machines  in  which  the  parts  are  worn  are  especially  liable 


The  Paraffin  Method  47 

to  this  defect.    It  may  be  remedied  to  some  extent  by  tightening  up  the 
parts  of  the  machine. 

b)  The  object  may  be  too  hard  for  the  knife  to  cut  and,  as  a  consequence, 
the  edge  of  the  knife  springs.    When  tough  or  hard  objects  must  be  cut, 
use  an  old  microtome  knife  or  a  sectioning  razor.    See  if  there  is  not  some 
means  of  softening  such  a  tissue  without  obscuring  the  microscopical  struc- 
tures sought. 

c)  Either  too  great  or  too  little  tilt  may  cause  the  defect  (step  13,  p.  40). 

d)  See  that  the  disk  bearing  the  object  is  securely  clamped  in  the  machine. 

8.  The  Object  Crumbles  or  Drops  Out  of  the  Paraffin  as  Cut. — It  has 
probably  been  insufficiently  penetrated  by  paraffin.    Some  of  the  following 
precautions  may  prevent  the  defect:    (1)  Leave  the  object  in  the  paraffin 
bath  longer.     (2)  See  that  it  is  entirely  free  from  the  dealcoholizing  fluid 
before  placing  it  into  the  melted  paraffin.    Objects  which  have  been  immersed 
in  cedar  oil  are  particularly  subject  to  this  defect.    For  this  reason  xylol  is 
better  than  cedar  oil  for  dealcoholization  in  general  work.     (3)  If  the  object 
is  impervious  to  paraffin  or  very  friable,  as  are  many  ova,  some  other  method 
must  be  tried.    Consult  memoranda  8  and  9;  see  also  the  celloidin  method 
(chap,  vii)  or  the  combination  celloidin-paraffin  method  (memorandum  8, 
p.  64). 

9.  The  Ribbon  Twists  or  Curls  About  or  Clings  Closely  to  the  Side  of  the 
Knife. — This  is  due  to  the  electrification  of  sections.     If  the  fault  is  excessive, 
it  is  best  to  postpone  the  cutting  until  the  atmospheric  conditions  have 
changed.    The  difficulty  may  be  minimized  by  using  a  drum  ribbon-carrier 
(see  step  15,  p.  40). 

10.  The  Cut  Section  Catches  On  and  Clings  to  the  Block  as  it  returns 
instead  of  remaining  on  the  knife.    Probably  the  knife  is  dull  or  its  edge  is 
dirty;   the  tilt  (step  13,  p.  40)  is  insufficient;   the  paraffin  is  too  soft,  or 
the  room  temperature  is  too  high. 

11.  A  Simple  Cooler  for  use  with  the  microtome,  which  facilitates  the 
preparation  of  thin  paraffin  sections  and  is  especially  useful  in  a  laboratory 
where  the  temperature  is  high,  is  described  by  Grave  and  Glaser  in  the 
Biological  Bulletin  for  September,  1910.    The  apparatus  "is  essentially  a 
hollow  truncated  pyramid,  open  at  both  ends,  and  suspended  in  an  inverted 
position  from  a  standard,  so  adjusted  that  the  lower  end  of  the  shoot  is  at 
a  convenient  distance  above  the  knife.    At  the  upper  end  of  the  inverted 
pyramid,  and  surrounded  by  it,  is  a  tray  whose  dimensions  are  less  than  those 
of  the  base  of  the  shoot.    This  tray  is  filled  with  crushed  ice,  and  from  one 
corner  of  it  a  drain  leads  the  water  to  the  escape  from  the  lower  end  of  the 
air-channel.    At  that  point  a  rubber  tube  connects  the  pipe  with  a  suitable 
receptacle."     Grave  and  Glaser  recommend  the  following  as  a  convenient 
size:  base,  12 -5X8 -7  in.;  truncated  apex,  6-1X2-1  in.;  measurements  of 
ice-tray,  8  -8X3  -3  in. 


CHAPTER  VI 

THE  PARAFFIN  METHOD:  STAINING  AND  MOUNTING 
I.    STAINING  WITH  HEMATOXYLIN 

Place  enough  of  the  following  reagents  in  tall  stender  dishes 
or  Coplin  staining-jars  to  cover  the  slides  lengthwise,  up  beyond 
the  sections  affixed  to  them:  xylol,  absolute,  95,  70,  50,  35  per  cent 
alcohols  respectively,  clear  water,  acid  alcohol,  and,  for  washing  out 
the  acid  alcohol  in  the  case  of  hematoxylin  preparations,  a  separate 
jar  of  70  per  cent  alcohol  to  which  a  few  drops  of  a  0.1  per  cent 
aqueous  solution  of  bicarbonate  of  soda  has  been  added.  Arrange 
these  reagents  in  a  row  in  the  order  named  with  the  exception  of  the 
acid  alcohol  and  its  accompanying  alkaline  alcohol  wash  of  70  per 
cent  alcohol,  which  may  be  placed  immediately  back  of  the  ordinary 
70  per  cent  alcohol.  Put  a  little  vaseline  along  the  upper  edges  of 
the  jars  containing  absolute  alcohol  and  xylol  and  press  the  cover 
down  tightly  to  prevent  evaporation  or  the  entrance  of  moisture. 

In  like  rrfanner  place  in  Coplin  staining-jars  (tall  stenders  will 
answer)  a  supply  of  Delafield's  hematoxylin  diluted  one-half  with 
distilled  water,  eosin,  Lyons  blue,  alum-cochineal,  Congo  red,  ^and 
solutions  A  and  B  for  the  iron-hematoxylin  method.  Arrange  these 
stains  in  a  row  back  of  the  alcohol  series. 

1.  Remove  the  paraffin  from  the  sections  of  intestine  (see  last 
lesson)  by  placing  the  slides  in  xylol  (turpentine  will  answer)  for 
10  or  15  minutes.     The  process  may  be  hastened  by  first  gently 
warming  the  slide  until  the  paraffin  begins  to  melt. 

2.  Remove  the  xylol  from  the  sections  by  transferring  the  slides 
to  absolute  alcohol  for  1  minute. 

3.  Pass  the  slides  through  the  alcohols  (95,  70,  50,  and  35  per 
cent),  leaving  them  for  a  half-minute  in  each. 

4.  Remove  to  Delafield's  hematoxylin  for  10  to  30  minutes  or 
until  stained  a  pronounced  blue. 

5.  Wash  in  water  for  \iminutes. 

48 


The  Paraffin  Method  49 

6.  Pass  the  slides  up  through  the  series  of  alcohols  to  70  per 
cent,  leaving  them  about  half  a  minute  in  each  alcohol. 

7.  Dip  each  slide  for  from  30  seconds  to  5  minutes  into  the  acid 
alcohol  until  the  sections  are  of  a  reddish  hue,  then  rinse  tiiem  in 
70  per  cent  alkaline  alcohol  until  the  blue  color  is  restored.     This 
last  alcohol  must  be  kept  very  slightly  alkaline  through  the  occa- 
sional addition  of  a  few  drops  of  a  0 . 1  per  cent  solution  of  bicarbonate 
of  soda  (see  memorandum  10,  p.  55).     The  alkaline  alcohol  may  be 
omitted  when  other  than  hematoxylin  stains  are  used,  as  its  purpose 
is  merely  to  restore  the  blue  color  of  the  latter. 

8.  Pass  the  slides  through  95  per  cent  alcohol   (3  minutes), 
absolute  alcohol  (5  minutes),  into  xylol  for  10  minutes  or  until  clear 
(see  memorandum  3,  p.  55). 

9.  Carefully  drain  off  all  excess  of  the  clearer  from  a  slide,  wipe 
the  under  side,  and  lay  it  down  flat  with  the  sections  uppermost. 
Put  a  few  drops  of  thin  balsam  on  the  sections  near  one  end.     Take 
up  a  clean  cover-glass  and,  holding  it  by  the  edges  between  the 
thumb  and  first  finger  of  one  hand,  lower  it  upon  the  balsam  by 
bringing  one  end  into  contact  with  the  slide  near  the  balsam  and 
supporting  the  other  end  by  means  of  a  needle  held  in  the  free  hand. 
Lower  the  cover  slowly  so  that  as  the  balsam  spreads,  no  air  bubbles 
will  be  inclosed  under  the  glass.     If  a  slide  is  tilted  a  little  and 
allowed  to  remain  in  that  position  small  bubbles  will  frequently 
work  out  unaided.     They  may  sometimes  be  removed  by  pressing 
gently  above  them  with  the  handle  of  a  needle  and  gradually  working 
them  to  the  edge  of  the  cover-glass.     Keep  the  slide  in  a  horizontal 
position  until  the  balsam  hardens. 

CAUTION. — Do  not  allow  the  sections  to  become  dry  before  adding 
the  balsam  and  cover. 

10.  Attach  the  permanent  label.     It  should  contain  at  least  the 
following  data:  the  number  of  the  record  card  (p.  5);  the  name  of 
the  tissue;,  the  kind  of  section  (plane  of  section,  thickness,  etc.); 
if  one  of  a  series,  the  number  of  the  slide  in  the  series  and  the  number 
of  the  first  and  last  section  on  the  slide;  the  date,  and  if  desired  the 
name  or  the  initials  of  the  preparator.     It  is  well  to  add  the  thickness 
of  the  cover-glass  (see  also  memorandum  23,  p.  58). 


50  Animal  Micrology 

It  is  best  to  have  the  label  on  the  left  end  of  the  slide,  as  it  will 
then  not  be  in  position  to  obscure  the  scale  of  the  detachable  mechani- 
cal stage  so  widely  used  on  microscopes  today. 

NOTE. — Prepare  four  slides  each  of  the  other  objects  which  have  been 
imbedded.  Stain  and  mount  one  of  each  kind  as  you  did  the  intestine,  and  also 
one  of  each  kind  in  the  same  way,  only  substitute  alum-cochineal  for  the  hema- 
toxylin.  The  alum-cochineal  may  require  12  hours  or  more  for  staining.  Pre- 
serve the  others  for  double  stairiing. 

As  time  permits,  prepare  and  section  the  other  tissues  which  were  fixed  in 
alcohol  and  in  Bouin's  fluid.  After  you  have  had  the  preliminary  practice  in 
double  staining,  stain  and  mount  these  as  you  prefer. 

II.     DOUBLE  STAINING  IN  HEMATOXYLIN  AND  EOSIN 

1.  Proceed  according  to  the  regular  schedule  with  one  each  of 
the  slides  reserved  above,  and  stain  in  Delafield's  hematoxylin. 

2.  Wash  the  sections  in  water,  and  proceed  farther  according 
to  the  regular  schedule  to  95  per  cent  alcohol. 

3.  Transfer  the  slide  to  the  eosin  stain  for  30  to  60  seconds,  and 
after  rinsing  again  in  95  per  cent  alcohol,  place  it  in  absolute  alcohol. 

4.  Clear  in  xylol  and  mount  in  balsam. 

NOTE. — The  sections  should  show  both  the  blue  stain  (in  nuclei)  and  the 
red  stain  (in  cytoplasm)  when  examined  under  the  microscope.  If  either  is  too 
dense  or  too  light,  make  a  note  of  the  fact  and  vary  the  time  accordingly  when 
staining  other  sections  by  this  method. 

m.    DOUBLE  STAINING  IN  COCHINEAL  AND  LYONS  BLUE 

1.  Pass  the  remaining  reserved  slides  through  xylol  and  the 
alcohols,  descending  to  35  per  cent  alcohol. 

2.  Stain  in  alum-cochineal  for  from  6  to  12  hours,  or  until  the 
sections  are  well  colored. 

3.  Rinse  in  water  or  35  per  cent  alcohol,  and  pass  the  sections 
up  through  the  alcohols  to  95  per  cent.     If  the  sections  are  deeply 
stained,  however,  remove  the  excess  of  stain  with  acid  alcohol  (a  few 
seconds)  when  the  sections  are  in  70  per  cent  alcohol. 

4.  Stain  for  10  to  20  seconds  in  Lyons  blue.     It  is  very  easy  to 
overstain  with  this  dye. 

5.  Rinse  in  95  per  cent  alcohol,  and  transfer  the  sections  to  abso- 
lute alcohol  (5  minutes),  clear  in  xylol,  and  mount  in  balsam. 


The  Paraffin  Method  51 

IV.    STAINING  WITH  HEIDENHAIN'S  IRON-HEMATOXYLIN 

This  stain  is  very  valuable  in  the  study  of  cell  division  and  in 
determining  the  finer  structure  of  the  nucleus.  The  iron-alum  acts 
as  a  mordant,  preparing  the  tissue  for  the  action  of  the  hematoxylin. 

1.  Prepare  two  sets  of  sections  of  intestine,  testis  or  ovary, 
bladder,  pancreas,  and  stomach.     The  sections  should  not  be  over 
6  or  7  microns  in  thickness.     Preserve  one  set  for  double  staining. 

2.  Pass  the  other  set  through  xylol,  absolute  alcohol,  95  per  cent 
alcohol,  and  thence  directly  into  water. 

3.  Transfer  from  water  to  the  iron-alum,  and  allow  this  solution 
to  act  for  from  30  minutes  to  1  hour. 

4.  Rinse  in  water  5  minutes. 

5.  Stain  in  the  0.5  per  cent  hematoxylin  1  hour.     If  a  trace  of 
the  iron-alum  remains  in  the  sections  the  hematoxylin  will  turn  black. 
This,  however,  does  not  impair  its  power  of  staining. 

6.  Rinse  in  water  5  minutes. 

7.  Place  the  sections  into  iron-alum  again,  which  will  now  extract 
the  excess  of  stain.     The  time  required  for  proper  differentiation 
varies  with  the  kind  of  tissue  and  the  fixing  agent  that  has  been  used. 
From  10  to  30  minutes  is  usually  sufficient,  though  no  definite  time 
limit  can  be  set.     Remove  the  slide  from  the  iron-alum  from  time 
to  time  and  inspect  it.     When  the  sections  become  of  a  dull-grayish 
hue  the  decolorization  is  usually  sufficient.     If  very  accurate  results 
are  necessary,  the  slide  should  be  removed  from  the  iron-alum 
frequently,  rinsed  in  water,  and  examined  under  the  microscope. 
When  in  a  dividing  cell  the  chromosomes  become  sharply  defined, 
the  decolorization  should  be  stopped. 

8.  Wash  hi  running  water  for  20  minutes  or  in  several  changes 
of  water  for  2  hours.     If  any  of  the  iron-alum  is  left  in  the  sections 
the  color  will  fade  later. 

9.  Wipe  off  the  excess  of  water,  transfer  the  slide  to  95  per  cent 
alcohol,  followed  by  absolute  alcohol  and  xylol. 

10.  Mount  in  balsam. 

NOTE. — Iron-hematoxylin  is  perhaps  the  one  most  important  stain  in  use 
today.  The  student  should  practice  the  method  until  he  has  mastered  it.  It 
is  better  though  not  absolutely  essential  that  the  stain  be  "ripe." 


52  Animal  Micrology 

For  demonstration  of  centrosomes  and  finer  cytological  details,  the  time  of 
staining  may  require  to  be  lengthened.  In  my  own  cytological  work  I  find  that 
2  to  4  hours  in  iron-alum  followed,  after  rinsing  in  water,  by  hematoxylin  for 
12  to  24  hours,  yields  better  results  than  does  the  shorter  method. 

V.    IRON-HEMATOXYLIN  WITH  OTHER  STAINS 

Use  the  sections  which  were  reserved  for  this  method.  The 
method  is  identical  with  the  one  just  outlined,  except  that  between 
step  8  and  step  9  the  following  directions  should  be  inserted:  8a, 
transfer  the  sections  from  water  to  Congo  red  for  a  minute,  or  to 
orange  G  for  2  hours,  then  wash  them  in  water  and  proceed  to 
step  9. 

NOTE. — Before  proceeding  farther,  kill  a  female  cat  or  rabbit  to  secure  tissues 
for  the  celloidin  method  and  to  correct  failures  in  the  paraffin  method.  In  addition 
to  the  tissues  specified  before,  prepare  (fix  in  Zenker's  fluid)  pieces  of  tendon,  carti- 
lage, spleen,  lymph  gland,  pancreas,  and  salivary  glands.  (If  the  reagents  are  at 
hand  and  time  permits,  the  student,  indeed,  might  advantageously  prepare  a  number 
of  tissues  according  to  the  methods  indicated  in  Appendix  C.)  Fix  the  ovary  in 
Bourn's  fluid,  and  reserve  it  for  the  paraffin  method  for  delicate  objects.  Fix  parts 
of  the  brain  and  cord  in  Zenker's  and  in  formalin  as  previously  indicated,  and 
place  bits  of  muscle  in  which  nerves  terminate  plentifully  (e.g.,  intercostals)  in 
formalin.  Larger  pieces  (up  to  2  cm.}  may  be  used  of  such  tissues  as  are  to  be 
imbedded  in  celloidin.  Bear  in  mind  that  the  larger  the  tissue  the  longer  must  it 
be  left  in  the  different  reagents.  Select  the  necessary  parts  of  the  digestive  tract  to 
prepare  longitudinal  sections  in  celloidin  from  esophagus  to  stomach  and  from 
stomach  to  intestine.  As  soon  as  possible  begin  the  preliminary  steps  in  the  celloidin 
method  (chap,  vii)  so  that  there  may  be  no  loss  of  time.  Prepare  a  piece  of  intestine 
for  staining  in  bulk  (see  vi).  It  should  be  placed  in  the  stain  after  thoroughly 
washing  out  the  fixing  reagent.  Preserve  parts  of  it  to  cut  in  celloidin.  Remove 
the  lower  jaw  and  prepare  it  for  decalcification  of  teeth  (as  indicated  in  chap.  xi). 
Likewise  prepare  pieces  of  femur  and  of  tar  sal  bone  for  sectioning  (chap.  xi). 

VL    STAINING  IN  BULK  BEFORE  SECTIONING 

It  is  sometimes  desirable  to  stain  objects  before  sectioning. 
The  method  is  a  slow  one,  and  requires  stains  which  penetrate  evenly 
and  thoroughly.  Various  preparations  of  carmine  and  cochineal 
give  the  best  satisfaction,  although  several  hematoxylin  stains  are 
also  frequently  used  in  this  way.  It  is  best  to  stain  immediately 
after  fixing  and  washing  out,  before  the  object  has  been  carried  into 
higher  alcohols.  In  general,  it  is  advisable  to  section  tissues  and 


The  Paraffin  Method  53 

stain  on  the  slides,  because  the  staining  can  be  controlled  more  effec- 
tually.    Use  the  piece  of  intestine  already  prepared  (see  note,  p.  52). 

1.  After  fixing  in  Zenker's  fluid  and  thoroughly  washing  according 
to  the  directions  on  p.  28,  place  the  tissue  in  Delafield's  hematoxylin 
for  24  hours. 

2.  Wash  in  35  per  cent  alcohol  for  5  minutes,  followed  by  50 
and  70  per  cent  alcohol,  20  minutes  each. 

3.  Decolorize  in  acid  alcohol  20  to  30  minutes,  or  until  the  color 
ceases  to  come  away  freely.     Restore  the  bluish-purple  tint  by  treat- 
ment with  alkaline  alcohol. 

4.  From  this  point  proceed  through  95  per  cent  alcohol,  absolute 
alcohol,  xylol,  and  imbedding,  sectioning,  and  mounting  precisely 
as  in  the  general  paraffin  method,  except  that  after  the  sections  have 
been  freed  from  paraffin  in  xylol,  do  not  mount  immediately  in  balsam, 
but  first  transfer  the  slide  back  into  absolute  alcohol,  and  thoroughly 
wash  it  in  order  to  remove  the  glycerin  from  the  fixative  and  so 
prevent  cloudiness  of  the  final  mount.     From  alcohol  the  slide  is 
passed  through  xylol,  or  carbol-xylol,  and  mounted  in  the  usual  way. 

NOTE. — When  certain  of  the  carmines  or  hematoxylins  are  used  as  stains 
for  entire  objects,  the  preparations  usually  need  to  be  decolorized  with  acid 
alcohol.  This  may  be  deferred,  however,  until  after  the  objects  are  sectioned. 

VH.    PARAFFIN  METHOD  FOR  DELICATE  OBJECTS 

To  prevent  the  distortion  of  delicate  objects  which  are  to  be 
sectioned  in  paraffin,  the  transition  of  the  material  from  one  reagent 
to  the  other  must  be  very  gradual  and  the  heat  be  minimized.  Ob- 
serve the  following  modifications  of  the  general  method  and  prepare 
pieces  of  ovary  which  have  been  fixed  in  Bouin's  fluid. 

1.  Pass  the  object  in  the  usual  manner  up  through  the  series 
of  alcohols  to  absolute.     It  is  sometimes  necessary  to  use  a  more 
closely  graded  series  of  alcohols  if  the  object  be  very  delicate.     See 
also  the  "drop"  method  (memorandum  6,  p.  152). 

2.  From  the  absolute  alcohol  pass  to  a  mixture  of  absolute  alcohol 
two-thirds  and  chloroform  one-third;  gradually  add  more  chloroform 
until  at  the  end  of  an  hour  the  mixture  is  at  least  two-thirds  chloro- 
form. 


54  Animal  Micrology 

3.  Transfer  to  pure  chloroform  for  30  minutes. 

4.  Add  melted  paraffin  little  by  little  during  the  course  of  an 
hour  or  two  (24  hours  will  do  no  harm),  until  the  chloroform  will 
hold  no  more  in  solution. 

5.  Transfer  the  object  to  pure  melted  paraffin  in  a  small  vessel 
on  the  paraffin  oven  for  10  to  20  minutes,  changing  the  paraffin  once. 
Imbed  in  the  usual  way. 

6.  Cut  the  sections  about  7  microns  thick.     Mount  and  stain 
some  in  Delafield's  hematoxylin   and    eosin   and   others  in  iron- 
hematoxylin  and  Congo  red  or  orange  G,  according  to  the  directions 
already  given  for  these  methods. 

NOTE. — For  very  sensitive  objects  Schultz's  dehydrating  apparatus  (to  be 
obtained  from  dealers)  may  be  used.  It  consists  of  a  tube  within  a  tube,  each 
having  the  lower  end  covered  by  an  animal  membrane.  The  tubes  are  suspended 
in  the  neck  of  a  much  larger  bottle  which  contains  95  por  cent  alcohol.  The 
object  is  placed  in  the  inner  tube  and  both  tubes  are  filled  with  water.  When 
suspended  in  the  alcohol,  a  very  gradual  hardening  or  dehydration  of  the  object 
.takes  place  as  the  alcohol  slowly  diffuses  through  the  membrane.  Sometimes 
it  is  necessary  to  use  only  one  tube,  and  in  such  a  case  the  hardening  proceeds 
more  rapidly. 

VIII.    EUPARAL  AS  A  MOUNTING-  AND  PRESERVATION-MEDIUM 

This  reagent,  introduced  by  Gilson,  has  been  highly  extolled  by 
Various  workers  as  a  final  mounting-medium,  although  most  still 
prefer  balsam  or  damar  for  general  work.  Two  forms,  colorless  and 
green,  are  obtainable  from  Griibler.  The  green  is  used  only  with 
hematoxylin  stains,  which  it  intensifies.  One  of  the  merits  of  euparal 
is  that  delicate  tissues  may  be  mounted  in  it  directly  from  95  per  cent 
alcohol,  thus  avoiding  the  expense  and  the  risks  of  passage  through 
absolute  alcohol  and  the  essential  oil.  Another  value  lies  in  the  fact 
that,  because  of  its  lower  index  of  refraction  (1.483),  unstained  or 
faintly  stained  elements  which  are  invisible  in  balsam  (index,  1 . 535) 
are  rendered  visible.  It  thus  becomes  particularly  serviceable  in 
the  study  of  spindle  fibers  and  other  such  delicate  cytological  ele- 
ments. It  hardens  rapidly,  hence  preparations  can  be  used  within 
24  hours.  It  is  well  for  the  beginner  to  try  it  along  with  other 
methods. 


The  Paraffin  Method  55 

MEMORANDA 

1.  In  Passing  from  One  Liquid  to  Another,  one  corner  of  the  slide 
bearing  sections  should  first  be  touched  by  blotting  paper  to  remove  any 
excess  of  the  liquid  last  used.    This  is  especially  necessary  in  transferring 
from  absolute  alcohol  to  xylol,  or  from  95  per  cent  to  absolute  alcohol. 

2.  Sections  Once  Placed  in  Turpentine  or  Xylol  for  the  removal  of  par- 
affin must  never  in  any  subsequent  step  be  allowed  to  become  dry.    Par- 
ticular care  must  be  taken  to  prevent  sections  from  drying  out  after  removing 
them  from  xylol  to  mount  in  balsam  because  the  xylol  evaporates  rapidly. 
If  the  sections  become  dry  the  preparation  is  usually  rendered  valueless. 

3.  Xylol  Used  for  Removing  Paraffin  should  be  kept  in  a  jar  separate 
from  that  which  contains  xylol  for  clearing  before  mounting,  and  it  should  be 
changed  occasionally  because  it  tends  to  become  saturated  with  paraffin. 

4.  Sections  Not  over  10  Microns  Thick  may  be  plunged  directly  from 
95  per  cent  alcohol  into  an  aqueous  medium  and  vice  versa.     If  sections 
are  over  10  microns  thick  it  is  better  to  put  them  through  the  complete 
series  of  alcohols.    With  thick  sections  diffusion  is  less  rapid,  and  too  abrupt 
a  change  from  one  fluid  to  another  may  produce  distortions  or  wrench  the 
sections  loose  from  the  slide. 

5.  To  Avoid  Rubbing  Sections  off  the  Slide,  hold  the  slide  with  one  end 
toward  the  light  before  wiping  it  and  glance  obliquely  along  the  surface. 
The  shiny  side  is  the  one  to  wipe. 

6.  The  Series  of  Alcohols  and  Stains  ordinarily  may  be  used  a  number  of 
times  without  replenishing.    When  the  alcohols  become  very  much  dis- 
colored or  the  stains  cloudy,  they  should  be  renewed.    Alcohols  should 
not  be  used  too  often,  however,  as  they  soon  accumulate  particles  of  dirt 
which  settle  upon  the  sections  and  render  preparations  unsightly. 

7.  Absolute  Alcohol  must  be  kept  free  from  water.    It  may  be  tested 
from  time  to  time  by  mixing  a  few  drops  with  a  little  turpentine.     If  the 
mixture  appears  milky  the  alcohol  contains  a  harmful  amount  of  water 
and  should  be  renewed. 

8.  Two  Slides  Placed  Back  to  Back  can  be  handled  as  readily  as  a  single 
slide  in  passing  through  the  various  liquids.    Various  kinds  of  slide-baskets 
or  slide-holders,  which  enable  one  to  transfer  a  number  of  slides  through 
liquids  at  one  time,  may  be  obtained  from  dealers. 

9.  Gentle  Agitation  of  a  Slide  in  any  liquid  facilitates  the  action  of  the 
liquid.    Observe  this  precaution  especiaUy  with  absolute  alcohol. 

10.  For  Washing  Sections  after  Staining  in  Hematoxylin  tap  water  is 
preferable  to  distilled  water  because  it  is  usually  slightly  alkaline.    When 
acid  alcohol  is  used  to  decolorize  sections  stained  in  hematoxylin,  the  sections 
should  be  washed  in  70  per  cent  alcohol  rendered  alkaline  by  the  addition 
of  a  few  drops  of  0. 1  per  cent  solution  of  bicarbonate  of  soda.    The  alkali 


56  Animal  Micrology 

neutralizes  the  acid  and  restores  the  bluish-purple  color  to  the  section;  it 
also  renders  the  blue  color  more  permanent.  If  too  much  of  the  soda  is 
added  the  color  will  be  a  hazy  disagreeable  blue. 

11.  To  Obtain  a  More  Precise  Stain  with  Delafield's  hematoxylin,  it  is 
well  to  dilute  it  with  three  or  four  times  its  bulk  of  distilled  water.    The 
sections  must  be  left  in  this  solution  a  correspondingly  longer  time.    Sec- 
tions stained  in  this  way  may  not  require  treatment  with  acid  alcohol. 
Most  workers,  however,  prefer  to  overstain  and  decolorize. 

12.  The  Length  of  Time  Required  for  Staining  Different  Tissues  is 
exceedingly  variable.    Upon  removal  from  the  stain  after  rinsing,  if  the 
sections  are  insufficiently  colored,  put  them  back  into  the  stain  and  examine 
them  from  time  to  time  until  they  are  properly  stained  (30  minutes  to  24 
hours). 

13.  If  Objects  Refuse  to  Stain,  it  is  usually  due  to  one  of  the  following 
causes:   (a)  The  fixing  agent  has  not  been  sufficiently  washed  out.    This  is 
a  frequent  cause  of  poor  staining.     (6)  The  fixation  has  been  poor.    The 
success  of  a  preparation  depends  largely  upon  proper  fixation  in  most  cases, 
(c)  The  stain  is  at  fault.    Hematoxylin  will  not  stain  properly  until  ripe 
(see  "Hematoxylin,"  p.  9).    Many  stains,  especially  the  anilins,  deteriorate 
and  must  be  replaced,     (d)  Certain  stains  will  not  follow  some  fixing  agents. 
This  can  be  remedied  only  by  using  a  different  stain  or  by  fixing  tissues  in  a 
different  fluid.    The  hematoxylins  and  carmines  are  applicable  after  a  very 
large  variety  of  fixing  agents,     (e)  The  paraffin  has  been  insufficiently 
removed  from  the  sections.    This  may  be  corrected  by  dissolving  off  the 
cover-glass  in  xylol  and,  after  thoroughly  removing  all  paraffin,  restaining 
and  mounting  the  sections  again  in  the  ordinary  way. 

14.  Use  Only  Clean  Slides  and  Covers. — Always  grasp  a  slide  or  a 
cover  by  its  edges  to  avoid  soiling  its  surface.    All  cloudiness  (seen  by 
looking  through  the  glass  toward  some  dark  object)  must  be  removed.    For 
wiping  slides  and  covers,  a  piece  of  cloth  which  does  not  readily  form  lint 
should  be  used.    A  well-washed  linen  towel  is  good,  as  is  also  bleached 
cheesecloth  cut  into  pieces  the  size  of  a  handkerchief.     Slides  may  often 
be  cleaned  after  simply  dipping  them  into  alcohol  or  into  alcohol  followed  by 
water.    If  this  treatment  is  insufficient,  place  them  for  several  hours  into 
equal  parts  of  hydrochloric  acid  and  95  per  cent  alcohol,  keeping  them  well 
separated  so  that  the  liquid  may  act  on  the  entire  surface  of  each.    Then 
rinse  them  in  water  and  place  them  in  ether-alcohol.    It  is  well  to  keep  a 
stock  supply  of  such  slides  and  cover-glasses  in  ether-alcohol. 

To  clean  a  cover-glass,  grasp  it  by  the  edges  in  one  hand,  cover  the 
thumb  and  first  finger  of  the  other  hand  with  the  cleaning-cloth,  and  rub 
both  surfaces  of  the  glass  at  the  same  time.  To  avoid  breaking  the  coverv 
keep  the  thumb  and  finger  each  directly  opposite  the  other.  A  large  cover- 


The  Paraffin  Method  57 

glass  may  be  cleaned  by  rubbing  it  between  two  flat  blocks  which  have  been 
wrapped  with  cleaning-cloths. 

To  clean  slides  which  have  been  used,  if  balsam  mounts,  warm  and  place 
in  xylol  or  turpentine  to  dissolve  off  the  covers.  Put  the  slides  and  covers 
into  separate  glass  or  porcelain  vessels  and  leave  them  for  a  few  days  in  the 
following  cleaning  mixture: 

Potassium  bichromate 10  parts 

Hot  water „ 50  parts 

Sulphuric  acid 50  parts 

In  making,  add  the  acid  very  cautiously  after  the  bichromate  solution 
cools.  When  the  slides  are  freed  from  balsam,  wash  them  in  water,  rinse 
in  a  dilute  solution  of  caustic  soda,  again  in  water,  and  finally  place  them  in 
ether-alcohol  until  needed. 

15.  If  Sections  Appear  Milky  or  Hazy  under  a  medium  power  of  the 
microscope,  when  finally  mounted,  the  effect  is  probably  due  to  one  of  the 
following  causes :  (a)  The  clearer  is  poor  and  needs  replenishing  or  correcting. 
(6)  The  absolute  alcohol  contains  water  (see  memorandum  7,  p.  55) .     (c)  The 
cover  bore  moisture.    Passing  a  cover-glass  quickly  through  a  flame  before 
putting  it  on  to  the  object  will  remove  moisture,     (d)  The  acid  has  not  been 
entirely  removed  from  the  sections,     (e)  Too  much  albumen  fixative  has 
been  used,    (f)  The  glycerin  of  the  albumen  fixative  has  not  been  removed 
by  passing  sections  of  objects  stained  in  bulk  (see  vi,  4,  p.  53)  back  into  abso- 
lute alcohol  after  removing  paraffin  from  them. 

The  defect  may  be  remedied  frequently  by  dissolving  off  the  cover  in 
xylol  or  turpentine,  descending  through  the  series  of  reagents  to  the  point 
where  the  fault  lies,  correcting  and  ascending  again  according  to  the  regular 
method.  To  remove  water,  for  example,  it  is  only  necessary  to  go  back 
as  far  as  absolute  alcohol  which  has  a  great  affinity  for  water. 

16.  Dry  or  Dull-Looking  Areas  under  the  Cover-Glass  indicate  that 
the  sections  were  allowed  to  get  dry  after  the  removal  from  the  clearer,  or 
that  insufficient  balsam  was  applied. 

17.  Delicate  Black  Pins  in  sections  of  tissue  fixed  with  a  mercuric  fixer 
are  due  to  deposition  of  mercury.    Remove  with  iodized  alcohol  before 
staining. 

18.  Balsam  Which  Exudes  from  under  the  Cover  may  be  scraped  off  with 
an  old  knife  after  it  hardens.     Remove  the  last  traces  by  means  of  a  brush 
or  a  cloth  dipped  in  turpentine  or  xylol.    Balsam  may  be  removed  from  the 
surface  of  a  cover  by  means  of  a  brush  dipped  in  xylol. 

19.  If  Sections  Wash  off  the  Slide  the  defect  is  probably  due  to  one  of 
the  following  causes:  (a)  The  slide  was  soiled  or  oily.    Remedy  by  cleaning 
slides  thoroughly  (see  14,  p.  56).     (6)  The  albumen  fixative  is  too  old. 


58  Animal  Micrology 

(c)  The  transitions  in  the  alcohol  have  been  too  great.  This  is  true  some- 
times of  thick  sections,  (d)  The  paraffin  ribbon  was  not  thoroughly  spread 
when  mounted  (see  p.  41). 

Thick  sections  are  more  likely  to  come  off  the  slide  than  thin  ones.  To 
avoid  this,  the  sections  may  be  collodionized  by  placing  them,  after  the 
paraffin  has  been  removed,  in  a  thin  solution  of  collodion  or  celloidin  in 
ether-alcohol  (f  gram  in  100  c.c.  of  ether-alcohol;  see  4,  p.  8)  for  a  few 
minutes  and  then  transferring  them  to  70  per  cent  alcohol.  If  carmine 
dyes  are  to  be  used,  this  method  is  not  satisfactory,  as  carmine  stains 
collodion. 

20.  Flooding  Sections  with  the  Dye  by  means  of  a  pipette,  especially 
in  case  of  stains  which  act  rapidly  (e.g.,  eosin,  acid  fuchsin,  Lyons  blue, 
picric  acid,  etc.),  is  sometimes  more  convenient  than  immersing  the 
sections  in  a  jar  of  the  staining  fluid.  Small  bottles  with  combination 
rubber  stopper  and  pipette  (Fig.  31)  are  now  provided  for  this  purpose 
by  dealers. 

21.  Balsam  Mounts  in  Which  the  Stain  Has  Faded  may 
frequently  be  restained,  either  with  the  original  or  with  other 
stains.    All  that  is  necessary  is  to  dissolve  off  the  cover  in  xylol 
(2  to   3  days)  and  pass  the  preparation  down  through  the 
alcohols  to  the  stain  in  the  usual  manner. 

22.  Ink  for  Writing  on  Glass  (Hubbert,  Journal  of  Applied 
Microscopy,  V,  1680). — Mix  drop  by  drop  3  parts  of  a  13  per 
cent  alcoholic  solution  of  shellac  with  5  parts  of  a  13  per  cent 

FIG.  31         aqueous  solution  of  borax.     If  a  precipitate  forms,  heat  the  solu- 
Bottie         tion  until  it  clears.    Add  enough  methylen  blue  to  color  the 

mass  deep  blue. 

Professor  Robert  F.  Griggs  uses  common  water-glass  (an  aqueous  solu- 
tion of  sodium  silicate  or  potassium  silicate)  with  an  ordinary  steel  pen. 
After  marking,  the  slide  is  heated  until  the  water-glass  decomposes,  leaving 
behind  a  rough,  sandy  surface,  which  when  rubbed  away  shows  the  written 
characters  etched  on  the  slide. 

"Diamond  ink"  obtainable  from  Eimer  &  Amend  is  useful  for  writing 
on  glass.  When  not  in  use  it  is  kept  sealed  with  paraffin.  See  also  p.  1, 
" Carborundum  Points." 

23.  More  Detailed  Labeling  than  that  indicated  on  p.  49  is  sometimes 
desirable.     For  well-devised  schemes  see  Richard  E.  Scammon,  "A  Method 
of  Recording  Embryological  Material,"  Kansas  University  Science  Bulletin, 
IV,  No.  5  (March,  1907);  also  Robert  T.  Hance,  "A  System  for  Recording 
Cytological  Material,  Slides  and  Locations  on  the  Slides,"  Transactions  of 
the  American  Microscopical  Society,  XXXV,  No.  1  (January,  1916). 

24.  For  Orientation  of  Objects  in  the  Imbedding-Mass,  see  p.  126. 


CHAPTER  VII 
THE  CELLOIDIN  METHOD 

Use  the  tissues  which  were  prepared  (p.  52)  for  this  method, 
including  pieces  of  the  brain  and  spinal  cord  which  were  fixed  in 
Zenker's  fluid.  Reserve  a  piece  of  spleen  for  the  freezing  method 
(p.  67). 

1.  Fixing,   washing,   and  dehydrating  are  the  same  as  usual 
(chap.  iii).     If  the  object  is  in  70  per  cent  alcohol,  complete  the  dehy- 
dration by  using  successively  95  per  cent  and  absolute  alcohol.     It 
should  remain  in  the  absolute  alcohol  for  from  12  to  24  hours. 

2.  From  absolute  alcohol  transfer  the  object  to  equal  parts  of 
absolute  alcohol  and  ether  12  to  24  hours. 

3.  Next  to  thin  celloidin  (reagent  20,  p.  12)  for  from  36  hours 
to  several  days  or  weeks. 

4.  Thence  to  thick  celloidin  for  from  24  hours  to  several  days. 
It  may  be  left  for  weeks. 

Thorough  dehydration  and  thorough  infiltration  are  the  great 
essentials  for  success  with  the  celloidin  method.  Some  workers 
prefer  to  use  a  graded  series  of  celloidins,  such  as  1J,  3,  6,  and  8  per 
cent,  leaving  the  object  in  each  for  from  24  hours  to  several  days  or 
even  weeks. 

5.  Prepare  a. wooden  block  (see  memorandum  3,  p.  63)  in  such 
a  manner  that  it  will  have  surface  enough  to  accommodate  the  object, 
leaving  a  small  margin,  and  length  enough  to  be  readily  clamped  into 
the  carrier  on  the  microtome  (Fig.  32).     Dip  the  end  of  the  block 
to  which  the  object  is  to  be  attached  into  ether-alcohol  for  a  minute 
and  then  into  thick  celloidin.     Let  it  dry  so  that  later  air  bubbles 
will  not  work  up  out  of  the  wood  into  the  imbedding  mass. 

6.  Oil  one  side  of  a  strip  of  stiff  paper  by  rubbing  on  a  very  little 
vaseline,  and  wrap  it,  oiled  surface  in,  about  the  prepared  end  of  the 
block  in  such  a  way  that  it  will  project  beyond  the  end  of  the  block, 
forming  a  collar  high  enough  to.  extend  a  little  beyond  the  object 

59 


60 


Animal  Micrology 


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The  Celloidin  Method  61 

which  is  to  be  placed  within  it.     Tie  the  paper  in  place  by  means  of  a 
thread. 

7.  Pour  a  small  amount  of  thick  celloidin  into  the  paper  cup  thus 
formed  and  with  forceps  remove  the  piece  of  tissue  and  place  it  in 
celloidin.    Add  more  thick  celloidin  until  the  cup  is  full.     By  means 
of  needles  which  have  been  moistened  in  ether-alcohol  arrange  the 
object  so  that  it  will  be  cut  in  the  desired  plane. 

NOTE. — Instead  of  being  mounted  in  blocks,  objects  may,  after  saturation 
with  thick  celloidin,  be  placed  in  proper  position  in  a  glass  dish  and  covered  with 
thick  celloidin.  The  dish  is  then  loosely  covered  and  placed  under  a  bell-jar 
so  that  the  ether  will  gradually  evaporate,  leaving  the  mass,  in  a  day  or  two,  of 
proper  consistency  for  cutting.  Each  specimen  is  then  cut  out  in  a  block  of 
suitable  size  for  sectioning.  For  fastening  to  base,  see  memorandum  5,  p.  63. 

8.  Into  a  small  stender  dish  put  chloroform  to  the  depth  of  3  mm. 
When  a  film  has  formed  over  the  exposed  surface  of  the  celloidin 
place  it  in  the  chloroform  to  harden.     It  need  not  be  submerged. 
Keep  the  vessel  tightly  covered.     The  object  may  be  left  for  a  day 
or  two,  but  1  to  3  hours  usually  suffices. 

9.  Transfer  the  block  to  70-83  per  cent  alcohol,  where  it  may 
remain  indefinitely. 

10.  Make  a  careful  study  of  the  microtome  used  for  cutting 
celloidin  (Fig.  32). 

11.  Place  the  block  in  the  object-carrier  of  the  microtome  at 
the  proper  level  and  arrange  the  microtome  knife  obliquely,  so  that 
it  will  slice  through  the  object  with  a  long  drawing  cut  for  at  least 
half  the  length  of  the  blade.     If  the  obj  ect  is  oblong  it  is  advantageous 
to  have  the  long  diameter  parallel  to  the  edge  of  the  knife. 

12.  Keep  both  the  knife  and  the  object  flooded  with  70  per  cent 
alcohol,  preferably  from  an  overhanging  drop-bottle. 

13.  Draw  the  knife  through  the  object  with  a  straight  steady 
pull;  avoid  pulling  down  on  or  lifting  the  knife-carrier. 

14.  If  the  feed  is  not  automatic  push  the  knife  back  to  position 
always  before  turning  the  screw  which  raises  the  object.     Cut  the 
sections  about  15  or  20  microns  thick.     If  they  curl  they  are  best 
unrolled  on  the  surface  of  the  knife  with  a  camel's  hair  brush  just 
before  they  are  wholly  cut  free  from  the  block. 


62  Animal  Micrology 

15.  As  the  sections  are  cut,  transfer  them  by  means  of  a  small 
soft  brush  or  a  paper  spatula  to  a  flat  stender  or  a  watch-glass  con- 
taining 70  per  cent  alcohol. 

16.  Transfer  some  of  the  sections  through  50  and  35  per  cent 
alcohol,  2  minutes  each,  into  alum-cochineal  for  from  20  to  30  minutes, 
or  until  stained  (12  to  24  hours). 

17.  Wash  successively  in  35,  50,  and  70  per  cent  alcohols,  leaving 
the  sections  from  2  to  3  minutes  in  each. 

18.  Transfer  the  sections  to  95  per  cent  alcohol  for  3  to  5  minutes. 
Absolute  alcohol  is  not  to  be  used  with  celloidin  because  it  dissolves 
the  celloidin. 

19.  Clear  in  cedar  oil  or  beechwood  creosote  for  from  10  to  20 
minutes. 

20.  Mount  in  balsam  (see  step  9,  p.  49). 

STAINING  CELLOIDIN  SECTIONS  IN  HEMATOXYLIN  AND  EOSIN 

The  objects  are  killed,  fixed,  and  preserved  as  usual  in  70  per 
cent  alcohol,  and  sectioned  as  in  the  foregoing  method. 

1.  Fifty  and  35  per  cent  alcohol,  each  3  to  5  minutes. 

2.  Delafield's  hematoxylin,  20  to  30  minutes,  or  until  stained. 

3.  Water,  5  minutes. 

4.  Thirty-five,  50,  and  70  per  cent  alcohol,  each  3  to  5  minutes. 

5.  Acid  alcohol,  until  the  celloidin  which  surrounds  the  object 
shows  but  little  of  the  stain. 

6.  Seventy  per  cent  alcohol,  barely  alkaline  (see  memorandum 
10,  p.  55),  until  the  red  color  caused  by  the  acid  is  replaced  by  bluish 
purple. 

7.  Alcoholic  eosin,  30  seconds  to  1  minute. 

8.  Ninety-five  per  cent  alcohol,  2  to  5  minutes.     Clear  in  cedar 
oil  or  beechwood  creosote  and  mount  in  balsam. 

NOTE. — As  time  permits,  section  other  tissues  by  the  celloidin  method  and 
stain  as  above. 

MEMORANDA 

1.  If  Chloroform  Is  Not  at  Hand,  80  per  cent  alcohol  will  harden  the 
celloidin,  although  more  slowly. 

2.  The  Length  of  Time  that  objects  should  be  left  in  ether-alcohol  and 
the  celloidin  mixtures  depends  upon  the  size  and  density  of  the  objects. 


The  Celloidin  Method  63 

When  time  permits,  it  is  always  best  to  leave  them  several  days  or  even 
weeks  in  the  mixtures  of  celloidin.  For  large  objects  such  as  the  medulla 
of  a  large  brain  this  is  a.  necessity.  For  an  embryo  of  large  size  months 
may  be  requited. 

3.  Blocks  for  Celloidin  Mounting  may  be  of  white  pine,  glass,  vulcanized 
fiber,  or  even  a  very  hard  paraffin.    Cork  should  not  be  used  because  it  is 
liable  to  give  or  bend.    The  vulcanized  fiber  is  the  most  satisfactory.    It 
may  be  purchased  from  dealers  in  the  form  of  strips  which  may  easily  be 
sawn  to  the  necessary  dimension.    It  is  well  to  saw  several  parallel  cuts  into 
the  upper  edge  of  the  block  to  provide  points  of  attachment  for  the  celloidin. 

4.  Other  Clearers  may  be  substituted  for  cedar  oil  or  creosote.    One 
which  clears  from  95  per  cent  and  which  does  not  dissolve  celloidin  must  be 
chosen.    Other  good  clearers  are:  (1)  origanum  oil;  (2)  a  mixture  of  3  parts 
of  oil  of  thyme  and  1  part  of  castor  oil;   (3)  Eycleshymer's  clearing  fluid, 
which  is  a  mixture  of  equal  parts  of  bergamot  oil,  cedar  oil,  and  anhydrous 
carbolic  acid. 

5.  Imbedding  a  Number  of  Objects  in  one  mass  is  frequently  convenient. 
Fold  a  stiff  paper  into  a  box  of  the  proper  size  (step  7,  p.  37)  or  use  metal 
L's  (Fig.  30).    Pour  in  thick  celloidin,  put  the  objects  in  place,  and  orient 
them  properly  for  cutting.    Leave  a  space  of  about  8  mm.  between  adjacent 
objects.    Fill  the  box  with  thick  celloidin  and  set  it  in  a  dish  containing  a 
little  chloroform,  or  leave  it  in  80  per  cent  alcohol  to  harden.    When  ready 
to  proceed,  cut  the  large  blocks  into  smaller  ones  each  containing  a  piece  of 
tissue.    To  fasten  it  to  the  wood,  trim  the  small  celloidin  block  to  the  proper 
dimensions,  soften  for  a  few  minutes  in  ether-alcohol  the  side  to  be  attached, 
then  dip  it  into  thick  celloidin  and  apply  to  the  end  of  a  wooden  block  which 
likewise  has  been  dipped  into  the  ether-alcohol  and  the  thick  celloidin. 
Press  the  two  together  and  place  them  in  chloroform  or  80  per  cent  alcohol 
to  harden  (see  also  note  under  step  7,  p.  61). 

6.  Anilin  Dyes  are  usually  avoided  in  the  celloidin  method  because  they 
stain  the  celloidin  intensely  and  are  not  removed  in  subsequent  treatment. 
When  necessary,  however,  some  (e.g.,  eosin)  may  be  used.    Safranin,  for 
example,  may  be  removed  satisfactorily  from  the  celloidin  by  means  of  acid 
alcohol  without  extracting  all  the  stain  from  the  tissue.    If  anilin  dyes  have 
been  used,  it  is  sometimes  better  to  remove  the  celloidin  by  treating  the 
sections  with  absolute  alcohol  or  with  ether  before  the  final  clearing  and 
mounting  (see  memorandum  15,  p.  66). 

7.  Relative  Merits  of  the  Paraffin  and  the  Celloidin  Methods.— Celloidin 
is  good  for  large  objects,  for  brittle  or  friable  objects,  and  for  delicate  objects 
which  heat  would  injure.    It  does  not  require  .removal  from  the  tissues 
ordinarily,  hence  it  holds  delicate  structures  together  permanently.    Some 
tissues  are  not  rendered  so  hard  and  so  difficult  to  cut  as  in  paraffin.     How.- 
ever,  very  thin  sections  cannot  be  obtained  except  by  great  skill.    The 


64  Animal  Micrology 

method,  moreover,  is  extremely  slow.  The  paraffin  method  is  comparatively 
rapid,  serial  sections  may  be  cut  and  mounted  with  ease,  and  very  thin 
sections  may  be  obtained.  Large  objects  do  not  section  as  satisfactorily 
as  in  celloidin,  although  up  to  10  mm.  or  even  considerably  greater  diameter 
they  cut  readily.  The  rule  is  to  use  the  paraffin  method  when  you  can. 

8.  For  Brittle  Objects,  a  Combination  of  Celloidin  and  Paraffin  Infil- 
tration sometimes  proves  successful  (see,  however,  memorandum  9,  p.  43). 
The  method  is  too  tedious  for  ordinary  use,  although  it  must  sometimes  be 
resorted  to  with  friable  or  delicate  objects  such  as  eggs. 

According  to  Apathy's  method,  as  reported  by  Kornhauser  (Science, 
July  14,  1916),  fixed  material  is  dehydrated  as  usual,  finally  passing  through 
three  changes  of  absolute  alcohol  into  ether  and  alcohol,  where  it  is  left  5 
hours.  It  is  next  put  into  2  per  cent  celloidin  for  24  hours,  then  4  per  cent 
celloidin  for  24  hours,  and  ultimately  imbedded  in  4  per  cent  celloidin  and 
hardened  in  chloroform  vapor  for  12  hours.  The  block  is  then  quickly 
trimmed,  leaving  a  margin  beyond  the  object  of  a  few  millimeters  on  every 
side,  and  put  into  liquid  chloroform  for  12  hours.  It  is  next  transferred  to  an 
oil  mixture  made  up  by  weight  instead  of  volume  as  follows : 

Chloroform 4  parts 

Origanum  oil 4  parts 

Cedar-wood  oil 4  parts 

Absolute  alcohol 1  part 

Carbolic-acid  crystals 1  part 

Anhydrous  sodium  sulphate  should  be  kept  in  the  bottom  of  the  tube 
to  take  up  any  water  brought  in,  in  the  celloidin.  . 

The  block  must  remain  in  this  oil  mixture  until  it  clears  and  sinks;  this 
may  take  from  3  days  to  a  week.  It  is  next  washed  in  three  or  more  changes 
of  benzol  to  remove  oils  and  alcohol,  then  infiltrated  in  paraffin,  imbedded, 
sectioned,  and  mounted  in  the  usual  way.  In  subsequent  handling,  slides 
should  not  be  left  for  any  great  length  of  time  in  absolute  alcohol,  as  it  will 
dissolve  out  the  celloidin. 

9.  To  Transfer  Celloidin  Sections  from  the  Knife,  it  is  an  excellent  plan 
to  use  a  paper  spatula;  a  bit  of  postal  card  held  in  the  cleft  end  of  a  small 
stick  answers  very  well.     Press  the  paper  down  evenly  on  the  section  and 
then  slide  it  off  the  edge  of  the  knife.    The  section  adheres  to  the  paper.     In 
carrying  loose  sections  from  one  fluid  to  another  an  ordinary  section-lifter 
may  be  used,  or  a  glass  rod  around  which  the  section  is  allowed  to  curl 
answers  very  well. 

10.  Objects  Stained  in  Bulk  May  Be  Cleared  While  Yet  in  the  Block, 
then  sectioned,  and  mounted  without  passing  back  into  the  alcohols.    After 
the  block  of  celloidin  has  hardened  sufficiently  in  chloroform  it  is  transferred 


The  Celloidin  Method  .          65 

directly  to  the  clearer  (cedar  oil,  or  a  mixture  of  oil  of  thyme  3  parts  and 
castor  oil  1  part).  In  cutting  objects  thus  cleared  the  knife  must  be  flooded 
with  the  clearer  instead  of  alcohol.  Do  not  allow  the  sections  to  become  dry. 
If  it  is  desired  to  use  this  method  for  a  celloidin  block  which  has  already  been 
preserved  in  70  to  83  per  cent  alcohol,  the  block  must  pass  through.  95  per 
cent  alcohol  (1  to  2  hours)  before  it  is  placed  in  the  clearer. 

11.  Collodion  instead  of  Celloidin  is  used  by  some  workers.    Celloidin, 
in  fact,  is  a  patent  preparation  of  collodion,  which  is  a  solution  of  guncotton 
(pyroxylin)  in  ether  and  strong  alcohol.    Thin  and  thick  solutions  are 
employed  and  the  method  is  in  every  respect  similar  to  the  celloidin  method. 

12.  Fixing  Celloidin  Sections  to  the  Slide  is  accomplished  (1)  by  cover- 
ing the  sections,  when  mounted  in  proper  order,  with  a  strip  of  tissue  paper, 
which  is  then  bound  fast  by  wrapping  thread  around  it.    Lee  (Microtomist's 
Vade-Mecum,  7th  ed.,  p.  125)  recommends  (2)  the  albumen  method  for 
celloidin  sections  as  well  as  for  paraffin.     (3)  If  the  sections  on  the  slide  are 
carefully  flooded  with  95  per  cent  alcohol  two  or  three  times,  this  drained 
off  and  followed  by  a  small  amount  of  ether-alcohol  or  ether  fumes  until  the 
edges  of  the  sections  begin  to  soften  perceptibly  (10  to  20  seconds),  the 
sections  will  generally  adhere  to  the  slide  sufficiently  when  the  celloidin 
becomes  hard  again  upon  exposure  to  the  air  (30  seconds)  after  the  ether- 
alcohol  has  been  drained  off;  they  must  then  be  immersed  in  95  per  cent 
alcohol  before  any  further  steps  are  taken. 

13.  For  Serial  Sections  in  Celloidin  some  one  of  the  so-called  plating 
or  sheet  methods  will  be  found  most  satisfactory.    That  of  Linstaedt 
(Anatomical  Record,  November,  1912)  is  among  the  best.    It  is  a  modifica- 
tion of  the  celloidin  sheet  method  suggested  by  Huber  for  paraffin  sections 
and  of  Weigert's  method  for  serial  sections  in  celloidin. 

Plates  of  glass,  thoroughly  clean  and  of  suitable  size  (e.g.,  5X7  inches), 
are  coated  on  one  side  with  the  following  solution  and  allowed  to  dry: 

Saccharose 3  grams 

Dextrin 3  grams 

Distilled  water 100  c.c. 

To  which  as  a  preservative  is  added  a  bit  of  thymol. 

When  the  sugary  layer  is  thoroughly  dry,  coat  it  with  a  4  per  cent  solu- 
tion of  celluloid  in  acetone.  A  number  of  such  sheets  may  be  made  up  and 
kept  dried  if  desired. 

As  sections  are  cut,  place  them  in  the  desired  order  on  the  celluloid  sheets, 
moistening  from  tune  to  time  to  prevent  drying  out.  Frilling  may  be  pre- 
vented by  brief  treatment  with  absolute  alcohol.  When  the  plate  is  filled 
with  sections,  blot  with  a  smooth-surfaced  toilet-paper,  then,  in  order  to 
fix  the  sections  to  the  celluloid,  spray  by  means  of  an  atomizer  with  a  1  to  2 


G6  Animal  Micrology 

per  cent  solution  of  celloidin  in  ether  and  alcohol.  When  it  is  partially  dry 
immerse  the  plate  in  70  per  cent  alcohol,  then  in  water.  The  water  will 
dissolve  the  sugary  solution  and  the  celluloid  sheet  bearing  the  sections  will 
float  off.  The  sheet  may  be  preserved  indefinitely  in  70  or  80  per  cent 
alcohol,  or  stained,  cleared,  and  mounted  at  any  time  by  any  of  the  methods 
suitable  for  ordinary  celloidin  sections.  Strips  may  be  cut  and  mounted 
serially  on  properly  numbered  slides. 

For  another  "sheet"  method  see  Bohm,  Davidoff,  and  Huber,  A  Text- 
book of  Histology,  1910,  p.  40. 

14.  Gilson's  Rapid  Celloidin  Process  (Lee,   The  Microtomist's    Vade- 
Mecum,  7th  ed.,  p.  112)  is  valuable  under  some  circumstances  because  of 
the  great  saving  of  time.    After  dehydration  the  object  is  saturated  with 
ether  and  finally  placed  into  a  test-tube  containing  thin  celloidin.    The  lower 
end  of  the  tube  is  then  dipped  into  melted  paraffin  and  allowed  to  remain 
there  until  the  celloidin  solution  has  boiled  down  to  about  one-third  of  its 
original  volume.    The  mass  is  then  mounted  in  the  ordinary  way,  hardened 
for  an  hour  or  more  in  chloroform,  and  cleared  in  cedar  oil.    Sections  are 
cut  as  directed  under  memorandum  10  above. 

15.  Celloidin  May  Be  Removed  from  Sections  when  necessary  by  passing 
them  through  absolute  alcohol  into  ether  and  alcohol  or  oil  of  cloves  for  from 
5  to  10  minutes,  then  back  through  absolute  to  ordinary  alcohol. 

16.  For  Orientation  of  Objects  in  the  Celloidin  Mass  see  p.  126. 


CHAPTER  VIII 
THE  FREEZING  METHOD 

1.  Use  a  piece  of  spleen  which  has  been  properly  fixed  and  later 
preserved  in  70  per  cent  alcohol.     Transfer  it  through  50  and  35 
per  cent  alcohol  successively  to  water,  and  wash  it  for  12  hours  in 
running  water. 

2.  Place  it  into  a  gum  and  syrup  mass  for  24  hours  (a  saturated 
solution  of  loaf  sugar  in  30  c.c.  of  distilled  water,  added  to  50  c.c. 
of  gum  mucilage.     Prepare  a  supply  of  gum  mucilage  by  dissolving 
60  grams  of  best  gum  acacia  in  80  c.c.  of  distilled  water). 

3.  Examine  the  freezing  microtome  carefully  (Fig.  33). 

4.  Remove  the  gum  and  syrup  mixture  from  the  outside  of  the 
tissue  with  a  cloth,  put  a  little  gum  mucilage  (not  gum  and  syrup) 
on  the  freezing  disk  of  the  microtome,  and  place  the  tissue  in  it  in 
such  a  way  that  longitudinal  sections  through  the  hilum  may  be  cut. 
Surround  the  object  with  gum  mucilage  and  set  the  freezing  apparatus 
to  going.     If  carbon  dioxide  is  used,  open  the  valve  very  cautiously, 
and  let  only  a  small  quantity  of  the  gas  escape. 

NOTE. — Carbon  dioxide  is  commonly  used  for  charging  soda  water  and  beer. 
It  may  be  purchased  in  iron  cylinders  containing  about  20  pounds  of  the  liquefied 
gas.  The  cylinder,  when  empty,  is  exchanged  for  a  charged  one,  so  that  the 
purchaser  pays  only  for  the  contents.  The  Bardeen  microtome  (Fig.  33)  may  ' 
be  screwed  directly  upon  the  carbon-dioxide  cylinder  when  the  latter  is  in  a 
horizontal  position,  or,  if  desired,  the  cylinder  may  be  placed  vertically  and  the 
microtome  attached  by  means  of  an  L-shaped  piece  of  heavy  tubing.  This 
microtome  has  the  advantage  over  the  common  forms  of  freezing  microtomes 
of  wasting  less  gas  and  of  greater  freedom  from  clogging.  It  is  advantageous  to 
have  an  extra  long  handle  to  the  key  which  is  used  for  opening  the  escape  valve 
of  the  carbon-dioxide  cylinder. 

Small  tubes  of  compressed  carbon  dioxide,  sufficient  for  one  or  two  freezings, 
may  sometimes  be  obtained  from  stores  carrying  automobile  supplies.  These 
can  be  utilized  in  operations  where  immediate  diagnosis  by  means  of  sections,  is 
required. 

5.  As  soon  as  the  gum  is  frozen,  continue  to  add  more  until  the 
tissue  is  completely  covered  and  frozen. 

67 


68 


Animal  Micrology 


6.  Work  the  microtome  screw  with  one  hand  and  plane  off  sections 
(15  to  20  microns  thick)  with  the  other.     The  well-sharpened  blade 
of  a  carpenter's  plane  is  the  best  instrument  for  cutting.     It  must  be 
frequently  stropped. 

NOTEV— The  blade  should  be  mounted  in  a  short,  broad  handle,  which  may 
be  grasped  easily  and  firmly  with  one  hand.  In  cutting,  the  bevel  edge  of  the 
knife  should  set  squarely  on  the  glass  ways  of  the  microtome  so  that  the  handle 

of  the  knife  is  inclined  toward  the  operator 
at  about  an  angle  of  45  degrees  from  the 
perpendicular.  The  hand  guiding  the 
knife  should  be  firmly  supported  against 
the  chest  while  pressing  the  cutting  edge 
steadily  against  the  glass  ways  of  the 
microtome.  The  cutting  stroke  is  made 
by  bending  the  body  forward  from  the 
waist  and  thus  forcing  the  blade  squarely 
across  the  surface  of  the  tissue. 

The  blade  must  be  kept  cold  to 
prevent  sections  from  sticking  to 
it.  If  the  sections  fly  off  or  roll, 
the  tissue  is  probably  frozen  too 
hard.  The  same  defect  may  arise 
if  there  is  insufficient  syrup  in  the 
gum  with  which  the  tissue  has  been 
saturated.  To  correct,  let  the  tissue 
thaw  a  little,  and  if  it  is  still  at  fault, 
soak  it  again  in  a  mixture  which 
contains  a  greater  proportion  of 
syrup.  Work  rapidly,  so  as  to 
cut  sections  in  quick  succession. 
Several  sections  may  be  allowed  to 
collect  on  the  blade  before  they  need  be  removed. 

7.  Transfer  the  sections  to  distilled  water.     The  water  should  be 
changed  several  times  to  dissolve  out  the  gum.     Reserve  a  few  sec- 
tions in  water  for  later  use  (step  11,  p.  69). 

8.  Immerse  a  few  of  the  sections  for  10  to  30  minutes,  or  until 
stained,  in  Delafield's  hematoxylin,  then  wash  them  in  several  changes 
of  tap  water. 


FIG.  33. — Bardeen  Carbon-Dioxide 
Freezing  Microtome 

The  freezing  chamber  contains  a 
spiral  passage  through  which  the  ex- 
panding carbon'  dioxide  passes,  securing 
the  maximum  freezing  power.  The 
knife  slides  on  glass  guides.  The  finest 
feed  is  20  microns. 


The  Freezing  Method 


69 


9.  Transfer  the  sections  through  the  successive  grades  of  alcohol 
(decolorizing  with  acid  alcohol  if  necessary)  up  to  absolute  alcohol, 
leaving  them  2  minutes  in  each,  after  which  remove  them  to  xylol  for 
5  minutes  or  until  clear.     If  desired,  stain  with  eosin  (30  to  60 
seconds)  after  70  per  cent  alcohol. 

10.  Remove  one  or  two  of  the  best  to  a  slide,  drain  off  the  excess  of 
xylol,  add  a  few  drops  of  balsam  and  a  cover-glass  of  suitable  size,  and 
label. 


FIG.  34. — Ether  or  Rhigolene  Freezing  Attachment 

11.  Remove  the  sections  reserved  in  step  7  (p.  68)  to  a  test-tube 
containing  a  small  amount  of  water,  and  shake  the  test-tube  vigor- 
ously for  a  minute  or  two.  This  removes  the  lymphocytes  from  the 
sections,  and  exposes  the  reticular  connective  tissue  so  that  it  may 
be  examined.  Dehydrate  the  sections  and  mount  in  balsam. 

MEMORANDA 

1 .  Fresh  Tissues  Are  Frequently  Sectioned  by  the  freezing  method.  The 
tissue  may  be  transferred  directly  to  the  disk  of  a  microtome  without  previ- 
ous imbedding,  and  sectioned  after  freezing.  This  affords  a  ready  means 


70  Animal  Micrology 

of  rapidly  determining  the  nature  of  a  given  tissue,  and  is  very  serviceable, 
especially  to  the  pathologist.  The  principal  objection  is  that  crystals  of 
ice  form  in  the  cells  and  distort  them  badly.  This  is  avoided  when  syrup 
and  gum  are  used  for  imbedding. 

2.  Fixed  Tissues  may  be  sectioned  by  the  freezing  method  if  they  are 
first  washed  in  running  water  for  some  hours.    Even  tissue  fixed  in  formalin — 
perhaps  the  best  fixing  fluid  for  tissues  which  are  to  be  frozen — is  better  foi 
a  washing  of  at  least  30  minutes  before  being  frozen. 

3.  Sections  May  Be  Preserved  in  alcohol  in  the  usual  way  after  being 
cut  by  the  freezing  method.    All  trace  of  gum  should  be  washed  out  and 
the  sections  passed  through  the  grades  of  alcohol  to  83  per  cent,  where  they 
may  remain  indefinitely. 

4.  Sections  of  Fresh  Tissue  May  Be  Fixed  and  washed  out  after  cutting 
if  desired.    This  requires  but  little  time,  and  the  sections  will  take  stain 
much  more  satisfactorily  after  having  been  subjected  to  a  fixing  reagent. 

5.  Objects  Which  Alcohol  Would  Injure  may  be  sectioned  by  the  freezing 
method  and  mounted  in  aqueous  media. 

6.  Ether  or  Rhigolene  Is  Sometimes  Used  for  Freezing,  although  the 
method  is  more  expensive  and  less  satisfactory  on  the  whole  than  the  carbon- 
dioxide  method.    Fig.  34  shows  a  common  form  of  freezing  attachment  used 
for  either  of  these  liquids. 

7.  Organs  or  Parts  Varying  Greatly  in  Density  may  sometimes  be  cut 
more  successfully  by  the  freezing  than  by  any  other  infiltration  method. 

8.  To  Fix  Frozen  Sections  to  the  Slide,  after  treatment  with  absolute 
alcohol,  flow  a  little  three-fourths  of  1  per  cent  solution  of  celloidin  in  ether 
and  alcohol  over  the  sections  and  drain  off  at  once.    After  a  few  seconds 
of  exposure  to  the  air,  place  in  80  per  cent  alcohol  for  a  minute.    The  film 
of  celloidin  should  be  very  thin.    If  it  turns  white  upon  immersing  in  the 
alcohol,  the  original  solution  of  celloidin  was  too  thick;  it  should  be  thinned 
by  adding  more  ether  and  alcohol. 


CHAPTER  IX 

METALLIC  SUBSTANCES  FOR  COLOR  DIFFERENTIATION 

I.    A  GOLGI  METHOD  FOR  NERVE  CELLS  AND  THEIR 
RAMIFICATIONS 

The  Golgi  chrom-silver  method  is  one  widely  used  for  the  demon- 
stration of  nerve  cells  together  with  their  various  processes.  There 
are  many  modifications  of  the  method,  all  of  which  are  more  or  less 
inconstant  in  their  results.  In  a  successful  preparation  the  various 
cells  and  nerve  processes  are  not  equally  blackened,  a  fact  which 
allows  of  discrimination  between  the  different  elements.  Sometimes 
the  ganglion  cells  and  fibers  remain  unstained  while  the  neuroglia 
cells  are  impregnated,  or  occasionally  other  elements  than  nervous 
tissue  (e.g.,  blood  vessels)  are  affected. 

The  following  method  is  applied  to  material  preserved  in  10 
per  cent  formalin  and  is  a  so-called  "rapid  method." 

1.  From  the  brain  and  spinal  cord,  which  have  previously  (see 
note,  p.  52)  been  subdivided  and  placed  in  at  least  10  times  their 
volume  of  10  per  cent  formalin  (3  days  to  an  indefinite  time),  cut 
out  small  pieces  4  to  5  mm.  thick  from  the  region  desired  for  study 
and  transfer  them  to  a  vessel  containing  from  15  to  20  times  their 
volume  of  a  3 . 5  per  cent  aqueous  solution  of  potassium  bichromate. 
They  should  remain  in  this  solution  for  from  2  to  5  days.     Renew 
the  fluid  at  the  end  of  12  hours.     Keep  the  different  pieces  of  tissue 
in  separate  vessels  so  as  to  avoid  confusion. 

2.  For  impregnation,  transfer  the  tissues  to  a  silver-nitrate  solu- 
tion made  as  follows: 

Silver  nitrate  (crystals) 1.5    grams 

Distilled  water 200    c.c. 

Concentrated  formic  acid 1    drop 

3.  Rock  the  tissues  gently  in  a  small  amount  of  this  fluid  until 
the  brown  precipitate  of  silver  chromate  ceases  to  appear,  then 

71 


72  Animal  Micrology 

transfer  them  into  from  20  to  40  times  their  bulk  of  fresh  silver- 
nitrate  solution  and  leave  them  in  the  dark  for  from  3  to  6  days. 
Change  the  fluid  after  the  first  12  hours. 

4.  Transfer  a  few  of  the  brown  pieces  of  tissue  to  95  per  cent 
alcohol  for  half  an  hour,  renewing  it  once  or  twice  during  this  time. 
Leave  the  rest  of  the  tissue  in  the  silver-nitrate  solution  for  future 
use  in  case  the  first  attempt  proves  unsuccessful. 

5.  Remove  the  pieces  from  95  per  cent  to  absolute  alcohol  for 
20  minutes,  changing  the  latter  once.     Then  transfer  them  to  ether- 
alcohol  for  20  minutes. 

6.  Imbed  in  celloidin  without  waiting  for  infiltration  to  occur 
(thin  celloidin  30  minutes,  thick  celloidin   10  minutes).     Mount 
directly  on  a  block  and  harden  in  chloroform  for  20  minutes. 

7.  From  chloroform  transfer  directly  to  the  clearing  fluid  (e.g., 
cedar  oil),  and  as  soon  as  clear  (30  to  60  minutes)  cut  sections  50  to 
100  microns  thick,  but  keep  the  knife  flooded  with  the  clearing 
fluid  instead  of  alcohol.     Cut  sections  of  cortex  so  that  they  will  be 
perpendicular  to  the  surface  of  the  brain. 

8.  When  the  sections  are  thoroughly  cleared,  transfer  them  to 
a  slide  flooded  with  the  clearing  fluid,  select  such  as  prove  desirable 
upon  microscopic  inspection,  and  discard  the  remainder. 

9.  Replace  the  oil  with  xylol,  then  remove  the  xylol  by  pressing 
upon  the  sections  with  blotting  paper.     Add  enough  thick  Canada 
balsam  to  cover  the  sections. 

CAUTION. — Do  not  put  on  a  cover-glass;  moisture  must  evapo- 
rate from  the  section.  If  this  is  prevented,  the  metal  deposits  break 
up  and  the  sections  become  worthless. 

10.  Keep  the  preparations  level  and  put  them  away  in  a  dry 
place  free  from  dust.     If  the  balsam  runs  off  the  sections,  more 
balsam  must  be  added  at  once.     Do  not  attempt  to  examine  under 
a  high  power  until  the  balsam  is  thoroughly  hardened. 

MEMORANDA 

1.  A  Fuller  Account  of  the  Golgi  Methods  will  be  found  in  Hardesty's 
Neurological  Technique  (pp.  55-61),  or  in  Lee's  Microtomisfs  Vade-Mecum 
(pp.  419-37). 


Metallic  Substances  for  Color  Differentiation  73 

2.  An  Osmium-Bichromate  Mixture  is  frequently  used  instead  of  for- 
malin for  fixing  fresh  tissues.    To  85  parts  of  a  3.5  per  cent  solution  of 
potassium  bichromate  add  15  parts  of  a  1  per  cent  solution  of  osmic  acid. 
Small  pieces  (4  to  6  mm.  thick)  of  fresh  tissue  are  placed  hi  40  times  their 
volume  of  this  mixture  and  kept  in  the  dark  for  from  12  to  24  hours.    This 
fixing  fluid  is  then  replaced  by  a  3.5  per  cent  solution  of  potassium  bichro- 
mate, as  in  the  case  of  material  fixed  in  formalin  (see  above).    From  this 
point  the  method  js  identical  with  the  one  given  above. 

3.  The  Determination  of  the  Elements  That  Will  Be  Impregnated 
appears  to  depend  upon  the  length  of  time  the  tissue  is  left  in  the  3 . 5  per  cent 
solution  of  potassium  bichromate.    Hardesty  gives  the  following  lengths 
of  time  for  different  structures:  neuroglia,  2  to  3  days;  cortical  cells,  3  to 
4  days;  Purkinje  cells,  spinal  cord,  peripheral  ganglion  cells,  4  to  5  days; 
nerve  fibers  of  the  spinal  cord,  5  to  7  days.    Axones  are  impregnated  ordi- 
narily only  in  so  far  as  they  are  not  medullated. 

4.  Mounting  the  Sections  upon  a  Cover-Glass  is  preferred  by  some 
workers.    The  cover-slip  is  then  fastened  over  the  opening  of  a  perforated 
slide  with  the  section  downward. 

5.  For  Permanently  Mounting  Golgi  Preparations  under  a  Cover-Glass 
Huber  recommends  the  following  method:  The  sections  are  removed  from 
xylol  to  the  slide  and  the  xylol  then  removed  by  pressing  blotting  paper 
over  the  sections.    A  large  drop  of  xylol-balsam  is  then  quickly  applied  and 
the  slide  is  carefully  heated  over  a  flame  from  3  to  5  minutes.    A  large  cover- 
glass  is  warmed  and  put  in  place  before  the  balsam  cools. 

6.  The  Cox  Modification  of  Golgi's  corrosive-sublimate  method  is  widely 
used.    It  is  likely  to  impregnate  nearly  all  of  the  cells  in  the  section.    This 
may  prove  to  be  disadvantageous  rather  than  otherwise,  however,  where 
cells  are  numerous  and  close  together.    Small  pieces  of  nervous  tissue  are 
placed  for  from  1  month  in  summer  to  2  or  3  months  in  winter  in  the  following 
solution : 

Potassium  bichromate,  5  per  cent  solution 20  parts 

Corrosive  sublimate,  5  per  cent  solution 20  parts 

Distilled  water ,. 30  to  40  parts 

Simple  chromate  of  potassium,  5  per  cent  solution     16  parts 

The  later  treatment  is  the  same  as  for  ordinary  Golgi  prepa- 
rations. 

7.  Tracheae  of  Insects,  Bile  Capillaries,  and  Gland  Ducts  may  also  be 
studied  by  the  Golgi  chrom-silver  method.    A  bit  of  the  wing  muscle  of  a 
bumble  bee  is  a  good  object  in  which  to  demonstrate  the  finer  ramifications 
of  tracheae. 


74  Animal  Micrology 

H.    OTHER  SILVER-NITRATE  METHODS 

a)  For  Nerves  (after  Hardesty) 

1.  The  fresh  nerve,  or;  better,  a  spinal  nerve  root,  may  be  obtained 
from  a  frog  which  has  j  ust  been  killed.     Without  stretching  the  nerve, 
carefully  insert  beneath  it  the  end  of  a  strip  of  postal  card  or  similar 
card  which  has  been  trimmed  to  the  width  of  50  mm.     The  nerve 
when  cut  off  at  each  side  of  the  card  will  adhere  to  it  and  remain 
straight  and  at  approximately  normal  tension. 

2.  Clip  off  the  end  of  the  card  bearing  the  nerve  into  a  clean 
vial  which  contains  0 . 75  per  cent  aqueous  solution  of  silver  nitrate. 
Place  the  vial  in  the  dark  for  from  12  to  24  hours. 

3.  Transfer  the  nerve  to  pure  glycerin  on  a  slide  and  tease  the 
fibers  apart  thoroughly  under  the  dissecting  microscope. 

4.  Add  a  cover-glass  and  expose  the  fibers  to  sunlight  until  they 
become  brown  (30  minutes). 

5.  To  make  the  preparation  permanent,  take  off  the  cover  and 
remove  the  glycerin  by  means  of  filter  paper,  add  a  few  drops  of 
warm  glycerin-jelly  (p.  96),  put  on  a  clean  cover-glass,  and  press  it 
down.     Wipe  away  the  exuded  jelly,  and  when  the  preparation  has 
cooled  seal  the  cover  with  gold  size,  followed  by  Bell's  cement  (see 
steps  5  and  6,  p.  95). 

The  preparation  should  show  the  "  cross  of  Ranvier"  and  the 
" lines  of  Fromman." 

b)    For  the  Cornea 

1.  Quickly  rub  a  piece  of  silver  nitrate  over  the  cornea  of  an  eye 
which  has  been  removed  from  a  recently  killed  frog. 

2.  Slice  off  the  cornea  and  place  in  distilled  water.     Brush  the 
surface  with  a  camel's  hair  brush  to  remove  the  epithelium  (conjunc- 
tivum). 

3.  Expose  to  the  action  of  sunlight  or  strong  daylight  until  the 
tissue  turns  brown. 

4.  Wash  in  distilled  water  and  mount  in  glycerin,  or  mount  in 
balsam,  after  proper  dehydration. 

If  the  preparation  is  successful  the  cells  should  be  strongly 
outlined    by    the   precipitated   silver.     If  desired,   after   washing, 


Metallic  Substances  for  Color  Differentiation  75 

the  nuclei  of  such  cells  may  be  stained  in  hematoxylin  according 
to  the  usual  method. 

MEMORANDA 

1.  Fresh  Membranes  are  also  commonly  treated  with  silver  nitrate  to 
outline  cells.    The  membrane  should  be  stretched  over  some  smooth  sur- 
face, or,  better,  by  means  of  two  small  vulcanite  rings  which  fit  one  into  the 
other  in  such  a  way  as  to  stretch  bits  of  membrane  like  a  drum-head,  and 
hold  them  fast.    Such  stretched  membrane  is  first  washed  with  distilled 
water,  then  agitated  in  a  1 : 300  silver-nitrate  solution,  hi  direct  sunlight, 
until  it  darkens.    It  is  then  washed  in  distilled  water,  removed  from  the 
rings,  and  mounted  in  glycerin,  glycerin- jelly  (p.  96),  or,  after  dehydration, 
in  balsam.    If  preferred,  after  washing  it  may  be  stained  in  hematoxylin 
to  bring  out  the  nuclei,  and  then  mounted. 

2.  Cajal's  Method  for  Neurofibrils  is  widely  used.    Small  pieces  of 
nervous  tissue  are  fixed  in  formalin  for  6  hours,  washed  in  water  4  hours,  and 
transferred  to  40  per  cent  alcohol  for  6  hours.    They  are  next  kept  for  24 
hours  in  40  per  cent  alcohol  to  which  ammonia  has  been  added  hi  the  pro- 
portion of  5  drops  of  ammonium  hydrate  to  50  c.c.  of  the  alcohol.    The 
tissues  are  then  placed  in  an  incubator  in  a  1 . 5  per  cent  silver-nitrate  solu- 
tion and  kept  for  5  days  at  a  temperature  of  38°  C.    Next  they  are  placed 
in  a  mixture  of  100  parts  of  water,  15  parts  of  formalin,  and  1  part  of  pyro- 
gallic  acid  or  hydrochinon  for  24  hours,  after  which  they  are  ready  to  be 
passed  through  graded  alcohols  into  paraffin  or  celloidin  and  sectioned  hi 
the  usual  way. 

An  excellent  application  of  the  Cajal  method  to  serial  sections  has  been 
devised  by  Malone  (Anatomical  Record,  IX  [1915],  791),  who  also  describes 
how  to  obtain  satisfactory  Cajal  preparations  from  sections  previously 
stained  by  the  Nissl  method. 

3.  The  Pyridine-Silver  Method,  a  modification  of  the  Cajal  method, 
devised  as  a  differential  stain  for  non-medullated  nerve  fibers,  has  come  into 
wide  use  in  American  laboratories  in  the  study  of  various  other  problems. 
It  is  often  used  in  the  preparation  of  sections  of  spinal  ganglia,  sympathetic 
ganglia,  and  spinal  cord  and  is  the  most  reliable  of  the  silver  stains.    Rela- 
tively large  pieces  of  tissue  can  be  successfully  stained.    Hanson's  technique 
(American  Journal  of  Anatomy,  XII  [1911],  69)  is  as  follows:   "The  nerve 
or  ganglion  is  placed  in  100  per  cent  alcohol,  with  1  per  cent  ammonia  for 
48  hours  (95  per  cent  -alcohol  with  5  per  cent  ammonia  will  give  much  the 
same  results,  but  seems  more  likely  to  bring  out  the  neurilemma  nuclei). 
The  pieces  are  then  washed  for  from  |  to  3  minutes  (according  to  their  size) 
in  distilled  water  and  transferred  to  pyridine  for  24  hours,  after  which  they 
are  washed  in  many  changes  of  distilled  water  for  24  hours.    They  are  then 


76  Animal  Micrology 

placed  in  the  dark  for  3  days  in  a  2  per  cent  aqueous  solution  of  silver  nitrate 
at  35°  C.,  then  rinsed  in  distilled  water  and  placed  for  1  to  2  days  in  a  4  per 
cent  solution  of  pyrogallic  acid  in  5  per  cent  formalin.  Sections  are  made 
in  paraffin,  and  ffter  mounting  are  ready  for  examination." 

For  use  of  the  method  in  staining  and  sectioning  the  entire  head  of  a 
small  animal  or  embryo,  after  decalcification,  see  Huber  and  Guild,  Anatomical 
Record,  VII  (1913),  253  and  331.  For  a  discussion  of  the  method  with  bibli- 
ography, see  Ranson,  Review  of  Neurology  and  Psychiatry,  November,  1914. 

IH.    GOLD-CHLORIDE  METHOD  FOR  NERVE-ENDINGS 

1.  Trace  some  of  the  motor  nerves  of  a  reptile  or  mammal  to 
where  they  enter  the  muscles  (intercostals  are  best),  and  clip  out 
small  pieces  of  the  muscle.     Use  material  that  has  been  preserved 
in  10  per  cent  formalin  (see  note,  p.  52). 

2.  Place  the  bits  of  muscle  in  10  or  12  times  their  volume  of  a 
10  per  cent  solution  of  formic  acid  in  distilled  water  and  leave  them 
for  from  30  to  40  minutes. 

3.  Transfer  the  tissue  into  from  8  to  10  times  its  volume  of  a  1 
per  cent  solution  of  gold  chloride  in  distilled  water  for  from  30  to  40 
minutes.     Avoid  direct  sunlight.     The  muscle  should  become  yellow 
in  color. 

4.  Remove  the  tissue  without  washing  it  to  about  25  volumes  of 
a  2  per  cent  formic-acid  solution  and  keep  it  in  the  dark  until  it 
assumes  a  purple  color  (24  to  48  hours).     When  the  fibers  appear 
reddish  violet  in  color  the  reduction  has  gone  far  enough;   if  they 
show  a  decidedly  bluish  tinge  the  process  has  gone  too  far. 

5.  Wash  the  tissue  in  several  changes  of  distilled  water  for  an 
hour  and  transfer  a  small  piece  to  a  slide.     Tease  the  fibers  apart 
very  carefully  under  a  dissecting  lens.     Great  care  must  be  exercised 
to  avoid  tearing  the  nerve  fiber  from  its  endings.     Examine  from 
time  to  time  under  a  low  power  of  the  compound  microscope,  and 
when  a  nerve  fiber  with  its  termination  is  found,  carefully  separate 
it  as  much  as  possible  from  the  other  fibers. 

6.  Add  glycerin-jelly  and  a  cover-glass.     Seal  in  the  ordinary 
way  (p.  95). 

NOTE. — Tissues  may  be  dehydrated  in  the  ordinary  way  and  mounted  in 
balsam  or  imbedded  in  paraffin  or  celloidin  and  sectioned. 


CHAPTER  X 

ISOLATION  OF  HISTOLOGICAL  ELEMENTS.     MINUTE 
DISSECTIONS 

I.    ISOLATION 

A.  Dissociation  by  Means  of  Formaldehyde;  ciliated  and  co- 
lumnar epithelium. — 1.  Kill  a  frog  and  secure  the  hinder  part  of  the 
roof  of  the  mouth,  bits  of  the  brain,  and  a  small  piece  of  the  intestine. 
Slit  open  the  latter.     Leave  the  objects  for  24  hours  in  a  dissociating 
fluid  made  by  adding  0. 5  c.c.  of  formalin  to  250  c.c.  of  normal  saline 
solution. 

2.  Scrape  the  roof  of  the  mouth  after  removal  from  the  fluid 
and  mount  the  ciliated  cells  thus  obtained  on  a  slide.     Similarly 
remove  some  columnar  epithelium  from  the  internal  surface  of  the 
stomach  and  mount  on  another  slide. 

3.  Add  a  cover-glass  and  examine.     If  the  cells  cling  together 
in  clumps,  separate  them  by  drumming  gently  upon  the  cover-glass 
with  the  handle  of  a  needle. 

4.  Stain  by  placing  a  drop  of  alum-cochineal  on  the  slide  just  at 
the  edge  of  the  cover  and  applying  a  bit  of  filter  paper  to  the 
opposite  edge  of  the  cover.     The  filter  paper  absorbs  the  fluid  from 
under  the  cover  and  the  stain  replaces  it.     Keep  the  preparations 
under  a  bell-jar  or  other  cover  to  prevent   evaporation  of  the 
staining  fluid. 

5.  After  a  few  hours  replace  the  stain  by  glycerin  in  a  similar 
manner. 

6.  If  a  permanent  preparation  is  desired,  the  cover-glass  must 
be  sealed  (p.  95),  or,  after  staining,  the  tissue  must  be  dehydrated 
and  mounted  in  balsam  in  the  usual  manner. 

B.  Isolation  of  Muscle  Fibers  by  Maceration  and  Teasing. — 
1.  Place  small  fragments  of  voluntary  muscle,  of  the  root  of  the 
tongue,  and  of  heart  muscle  of  the  frog  into  separate  vials  containing 
MacCallum's  macerating  fluid  (reagent  89,  p.  238).     After  2  days 

77 


78  Animal  Micrology 

pour  off  the  fluid,  fill  the  vials  about  half  full  of  water,  and  separate 
the  fascicles  by  shaking  each  vial.  Further  isolate  the  fibers  by 
teasing. 

Teasing. — In  teasing,  the  important  thing  to  remember  is  that  the 
elements  of  the  tissue  are  to  be  separated,  not  broken  up.  Both  patience 
and  sharp  clean  needles  are  indispensable.  The  process  is  best  carried  on 
under  the  lens  of  a  dissecting  microscope  or  a  binocular  dissector,  although 
it  may  be  done  without  such  aid.  A  background  which  enables  the  tissue 
to  be  seen  distinctly  should  be  selected,  black  for  colorless  or  white  for  colored 
objects.  Black-and-white  porcelain  slabs  are  made  for  this  purpose  and  are 
very  convenient.  A  good  dissecting  microscope  has  attached  beneath  the 
stage  a  reversible  plate  one  side  of  which  is  black,  the  other  white.  Use  a 
small  piece  of  tissue  and  begin  teasing  at  one  end  of  it. 

2.  With  the  aid  of  a  dissecting  microscope  carefully  tease  out 
in  water  a  number  of  fibers.     Use  a  small  piece  and,  beginning  at 
one  end,  with  both  needles  separate  the  piece  along  its  entire  length 
into  two;  likewise  further  subdivide  these  until  the  ultimate  fibers 
are  isolated. 

3.  Transfer  some  of  the  fibers  through  the  alcohols  and  xylol 
and  mount  in  balsam.     Stain  others  in  alum-cochineal  for  some 
hours  and  mount  in  glycerin  as  above. 

C.  Maceration  by  Means  of  Hertwig's  Fluid  (Hydra.  Testis). — 
1.  The  solution  consists  of: 

0.05  per  cent  aqueous  solution  of  osmic  acid 1  part 

0.2    per  cent  acetic  acid 1  part 

Prepare  the  ingredients  for  this  mixture  by  diluting  the  stock 
solution  (1  per  cent)  in  each  case  with  distilled  water.  Make  a 
separate  0 . 1  per  cent  solution  of  acetic  acid  also. 

2.  Treat  a  hydra  with  the  osmic  and  acetic  acid  mixture  for  3 
minutes  and  then  transfer  it  to  the  0 . 1  per  cent  solution  of  acetic 
acid.    Wash  in  several  changes  of  this  fluid  to  remove  all  osmic  acid 
and  let  the  hydra  remain  in  the  acetic  acid  for  12  hours. 

3.  Wash  in  water,  stain  in  alum-cochineal  or  in  acid  carmine 
(reagent  38,  p.  222)  and  mount  in  glycerin  as  above.     If  the  cells 
are  not  sufficiently  separated,  gently  tap  on  the  cover-glass. 


Isolation  of  Histological  Elements  79 

4.  Submit  small  bits  of  the  testis  of  some  animal  to  the  same 
treatment.  Stain  with  methyl  green  (reagent  60,  p.  231)  or  acid 
carmine  (reagent  38,  p.  222). 

D.  Mall's  Differential  Method  for  Reticulum. — 1.  Cut  sections 
of  fresh  spleen  or  lymph  gland  40  to  80  microns  thick  by  the  freezing 
method  and  digest  for  24  hours  in  the  following  solution : 

Pancreatin  (Park,  Davis  &  Co.) 5  grams 

Bicarbonate  of  soda 10  grains 

Water 100  c.c. 

2.  Wash  thoroughly  in  water,  then,  in  order  to  remove  cellular 
debris,  shake  for  some  minutes  in  a  test-tube  half  full  of  water. 
Spread  out  on  a  slide  and  allow  to  dry. 

3.  Apply  a  few  drops  of  a  3 . 5  per  cent  solution  of  picric  acid  in 
11  per  cent  alcohol  and  allow  it  to  dry  on  the  preparation. 

4.  Stain  for  about  half  an  hour  in  a  10  per  cent  solution  of  acid 
f  uchsin  in  35  per  cent  alcohol. 

5.  Wash  in  the  picric-acid  solution  (step  3)  for  a  moment,  then 
pass  through  alcohol  and  xylol  and  mount  in  balsam. 

H.    MINUTE  DISSECTIONS 

A.  Alimentary  Canal  and  Nervous  System  of  Insects. — 1.  Carefully 
dissect  out  the  alimentary  canal  and  the  central  nervous  system  of  a  cock- 
roach with  the  aid  of  the  dissecting  microscope  or  lens.    Wash  each  by 
gently  flooding  it  with  distilled  water  from  a  pipette,  and  then  cover  it 
with  Bouin's  fluid  or  corrosive  sublimate  (reagent  14,  p.  212)  for  30  minutes. 

2.  Wash  in  several  changes  of  water  during  the  course  of  half  an  hour  and 
stain  for  40  minutes  or  more  in  borax-carmine. 

3.  Wash  in  50  per  cent  alcohol  and  decolorize  in  70  per  cent  acid  alcohol 
until  the  objects  become  bright  scarlet  in  color. 

4.  Wash  in  95  per  cent  alcohol  for  5  minutes  and  then  transfer  to  absolute 
alcohol  for  5  minutes,  xylol  or  turpentine  10  minutes,  and  mount  in  balsam. 
Apply  cover  and  label. 

B.  Gizzard  of  Cricket  or  Katydid. — Pull  off  the  head  of  a  cricket  or 
katydid.    The  gizzard  usually  remains  attached  to  the  head  part.    Cut 
it  open  lengthwise,  wash  out  the  contents  and  mount  as  above,  but  omit  the 
staining.    The  inside  should  be  turned  uppermost. 

C.  Sting  of  Wasp  or  Bee. — 1.  Place  a  wasp  or  bee  in  water,  cover  to 
keep  out  dust,  and  let  it  stand  for  two  or  three  days  until  the  smell  becomes 
unpleasant. 


80  Animal  Micrology 

2.  Wash  in  clear  water  and  squeeze  the  abdomen  gently  until  the  sting 
protrudes.    With  forceps  pull  it  out  carefully.    The  poison  gland  and  duct 
should  come  away  with  it. 

3.  Place  the  parts  removed  on  a  slide  and  under  a  lens  draw  the  sting 
out  of  its  sheath  by  means  of  a  small  needle  which  should  be  drawn  over  the 
outer  surface  of  the  sheath  from  the  base  to  the  apex  of  the  sting. 

4.  Stain  and  follow  out  the  same  subsequent  treatment  as  for  II,  A, 
above,  or  mount  without  staining.     It  is  advisable  to  compress  the  object 
between  two  slides  as  soon  as  the  acid  alcohol  is  washed  out.    The  slides 
•should  be  tied  together  and  left  in  95  per  cent  alcohol  several  hours.    Then 
proceed  in  the  ordinary  way. 

D.  Salivary  Gland  of  Cockroach  or  Cricket. — Let  the  animal  soak  in 
water  as  for  preparation  of  sting.    When  sufficiently  decayed  pull  off  the 
head  carefully  with  forceps.    The  esophagus,  the  salivary  glands,  and  crop 
usually  come  along  with  it.    Stain  and  mount  as  for  sting.    For  preparation 
of  fresh  salivary  gland  use  the  "salivary  gland  of  a  Chironomous  larva." 

E.  Mouth  Parts  of  Insect. — 1.  Place  the  head  of  a  bee  or  cockroach 
in  95  per  cent  alcohol  for  2  or  3  hours.    Transfer  to  absolute  alcohol  for 
30  minutes,  and  then  to  cedar  oil  for  30  minutes  to  an  hour. 

2.  Remove  the  head  to  a  slide  and  in  a  drop  of  the  oil  dissect  out  the 
mouth  parts.  Transfer  them  to  a  clean  slide,  remove  the  excess  of  oil,  and 
arrange  them  in  their  relative  positions  in  sufficient  balsam  to  hold  them  in 
place,  then  set  the  slide  aside  in  a  place  free  from  dust  until  the  balsam 
hardens  enough  to  keep  the  parts  from  shifting.  Make  any  necessary 
rearrangement.  Add  more  balsam  and  a  cover. 

MEMORANDA 

1.  The  Cover-Glass  May  Be  Supported  by  means  of  small  wax  feet, 
bits  of  broken  cover-glass,  or  fine  glass  threads  when  the  tissue  is  too  bulky 
to  allow  the  cover-glass  to  fit  down  closely  to  the  slide. 

2.  A  General  Rule  for  Dissociating  Tissues  is  to  use  small  pieces  of  the 
tissue  and  not  a  very  great  amount  of  the  fluid. 

3.  For  Minute  Dissections  clove  oil  is  often  a  convenient  medium.     It 
tends  to  form  very  convex  drops,  clears  well,  and  renders  the  object  brittle; 
any  or  all  of  which  properties  may  be  useful  in  such  dissections. 

4.  The  Fixation  of  Pieces  of  Macerated  Tissue  (e.g.,  macerated  epithe- 
lium) in  0 . 5  to  1  per  cent  osmic  acid  for  an  hour  or  so  often  proves  advan- 
tageous. 

5.  Congo  Glycerin  is  recommended  by  Gage  as  especially  good  for  iso- 
lated preparations,  particularly  nerve  cells.     It  is  made  by  dissolving  |  gram 
of  Congo  red  in  glycerin.     It  acts  both  as  a  stain  and  as  a  mounting-medium. 
The  preparation  may  be  sealed  (p.  95)  if  desired. 


CHAPTER  XI 
TOOTH,  BONE,  AND  OTHER  HARD  OBJECTS 

Sectioning  Decalcified  Tooth. — 1.  Kill  a  cat  and  remove  the 
lower  jaw  (p.  52).  With  a  fine  saw  cut  out  about  a  quarter  of  an 
inch  of  the  bone  bearing  a  tooth  (e.g.,  canine),  remove  as  much  of  the 
surrounding  tissue  as  possible,  and  place  the  object  in  Zenker's 
fluid  for  1  or  2  days.  Wash  thoroughly  in  water  and  place  in  alcohol 
for  at  least  24  hours.  Transfer  to  nitric-acid  decalcifying  fluid 
(reagent  10,  p.  9).  Use  a  relatively  large  quantity  of  the  fluid  and 
change  it  each  day  until  the  tooth  is  decalcified  (2  to  6  days) .  It  is 
sufficiently  soft  to  cut  when  a  needle  can  be  thrust  into  it  easily. 
Use  this  test  sparingly,  however,  as  it  injures  the  tissue. 

2.  Wash  it  in  repeated  changes  of  70  per  cent  alcohol  until  all 
trace  of  the  acid  is  removed. 

3.  Transfer  the  object  through  50  and  35  per  cent  alcohol  suc- 
cessively to  running  water  and  wash  for  24  hours. 

4.  Cut  sections  by  means  of  the  freezing  microtome  as  directed 
under  that  method  (p.  67).     If  a  freezing  microtome  is  not  available 
use  the  celloidin  method. 

5.  After  dissolving  out  all  of  the  gum  from  the  sections  in  distilled 
water,  stain  in  alum-cochineal  and  Lyon's  blue  (see  method,  p.  50). 
Dehydrate.     Remove  one  or  two  of  the  best  sections  (through  the 
center  of  the  tooth)  to  a  slide,  clear,  and  mount  in  the  usual  way  in 
balsam. 

6.  Stain  other  sections  in  1  per  cent  osmic  acid  for  24  hours  and 
mount  in  glycerin-jelly.     When  the  jelly  has  hardened,  seal  the  cover 
with  gold  size,  and  when  this  is  dry  add  a  thin  coat  of  Bell's  cement 
(see  p.  95).     If  preferred,  dehydrate  and  mount  in  balsam  instead  of 
glycerin-jelly. 

Sectioning  Decalcified  Bone. — Saw  out  a  short  piece  from  the 
femur  of  a  cat  (p.  52).  Prepare  transverse  sections  by  decalcifying 

81 


82  Animal  Micrology 

and  sectioning  in  the  same  manner  as  for  teeth.  Do  not  destroy 
the  periosteum.  Prepare  likewise  longitudinal  sections  of  a  tarsal 
bone. 

Sectioning  Bone  by  Grinding. — 1.  With  a  fine  saw  cut  a  thin 
transverse  section  of  the  femur  of  a  cat.  Let  it  macerate  in  water 
until  quite  clean,  then  dry  it  carefully. 

2.  Grind  the  disk  of  bone  between  two  hones,  keeping  the  hones 
parallel  in  order  to  avoid  wedge-shaped  sections.     The  section  is  not 
thin  enough  until  fine  print  can  readily  be  distinguished  through  it. 

3.  Wash  the  section  thoroughly  in  water,  transfer  it  to  absolute 
alcohol  for  10  minutes,  then  to  pure  ether  for  half  an  hour. 

4.  After  removal  from  the  ether,  clamp  it  between  two  slides 
by  means  of  a  string  or  a  rubber  band  and  let  it  dry  thoroughly. 

5.  Place  some  xylol-balsam  in  the  center  of  a  slide  and  heat  it 
for  a  few  minutes  to  drive  off  the  xylol,  then  press  the  section  of  bone 
down  firmly  into  it  and  put  on  the  cover-glass.     The  air  in  the  spaces 
of  the  bone  makes  them  stand  out  black.     The  balsam  should  not  be 
thin  enough  to  enter  these  spaces. 

MEMORANDA 

1.  Failure  to  Stain  Properly  is  due  ordinarily  to  insufficient  washing 
out  of  the  acid. 

2.  Teeth  and  Other  Hard  Objects  may  be  prepared  by  grinding  in  the 
same  way  as  bone. 

3.  For  Other  Decalcifying  Fluids  than  nitric  acid,  see  Appendix  B,  v. 


CHAPTER  XII 
INJECTION  OF  BLOOD  AND  LYMPH  VESSELS 

Red  Injection  Mass. — 1.  Rub  up  4  grams  of  carmine  thoroughly 
with  8  c.c.  of  distilled  water  in  a  mortar  and  add  ammonium  hydrate 
drop  by  drop  until  a  transparent  red  color  results. 

2.  After  quickly  washing  it  to  remove  dust,  etc.,  soak  10  grams  of 
best  French  gelatin  in  distilled  water  until  it  is  swollen  and  soft  (18 
hours),  then  remove  it  to  a  porcelain  evaporating-dish  and  melt  it  at 
a  temperature  of  about  45°  C. 

3.  While  the  gelatin  is  yet  fluid,  slowly  add  the  coloring  matter, 
stirring  constantly  until  a  homogeneous  mixture  is  obtained. 

4.  Before  the  mass  cools  add  also  some  25  per  cent  acetic-acid 
solution  drop  by  drop,  stirring  thoroughly  until  the  mass  becomes 
slightly  opaque  and  the  odor  of  ammonia  gives  place  to  a  faint  acid 
smell.     Watch  for  this  change  closely,  for  a  few  drops  too  much  of  the 
acid  will  spoil  the  entire  mass  by  precipitating  the  carmine.     If  the 
ammonia  is  not  completely  neutralized,  on  the  other  hand,  the  color- 
ing matter  will  diffuse  through  the  walls  of  the  injected  vessels 
and  stain  the  surrounding  tissues.     Walker  (American  Journal  of 
Anatomy  [1905],  p.  74)  makes  results  more  certain  by  mixing  1  part 
of  the  laboratory  ammonia  with  4  parts  of  distilled  water,  and  then 
determining  the  exact  amount  of  the  laboratory  acetic  acid  which 
will  neutralize  it.     Knowing  this,  it  is  easy  to  determine  the  total 
amount  of  acetic  acid  which  must  be  added  for  the  amount  of 
ammonia  which  has  been  used  in  any  quantity  of  the  gelatin  mass. 
Just  before  using,  the  mass  should  be  heated  and  strained  through 
clean  flannel  wrung  out  of  hot  water. 

With  a  large  animal  it  is  advisable  to  keep  animal  and  apparatus 
submerged  in  warm  normal  saline  during  the  operation  of  injection, 
but  with  a  small  animal  this  is  unnecessary  if  the  operator  works 
rapidly. 

83 


84  Animal  Micrology 

Blue  Injection  Mass. — Prepare  a  gelatin  mass  as  directed  above.  To 
the  warm  mass  add  sufficient  quantity  of  saturated  aqueous  solution  of 
Berlin  blue  to  give  the  desired  blue  color.  If  the  blue  does  not  dissolve, 
add  a  little  oxalic  acid  to  the  mixture.  The  blue  mass  need  not  be  made 
for  the  present  practical  exercise  unless  the  student  wishes  to  undertake  a 
double  injection  as  indicated  in  memorandum  2,  p.  87. 

Yellow  Injection  Mass. — Prepare  a  gelatin  vehicle  consisting  of  1  part 
of  gelatin  to  4  parts  of  distilled  water.  Take  equal  volumes  of  the  gelatin 
mass,  a  cold,  saturated  solution  of  bichromate  of  potassium,  and  a  cold, 
saturated  solution  of  lead  acetate.  Add  the  bichromate  solution  to  the 
gelatin  and  heat  almost  to  boiling;  then  add  slowly,  while  stirring,  the  solu- 
tion of  lead  acetate. 

INJECTING  WITH  A  SYRINGE;    SINGLE  INJECTION 

A  common  method  of  injection,  and  one  which  proves  satisfactory 
in  many  instances,  is  by  means  of  a  metal  or  glass  syringe.  Although 
not  as  desirable  in  the  main  as  the  method  of  continuous  air  pressure, 
many  good  injections  may  be  made  by  means  of  the  syringe.  The 
apparatus  consists  of  a  syringe  fitted  with  a  stop-cock  in  the  nozzle, 
and  a  separate  tube,  known  as  the  cannula,  which  fits  on  to  the  end 
of  the  nozzle.  The  syringes  are  made  in  different  sizes,  and  each  is 
provided  with  an  assortment  of  cannulae  to  fit  vessels  of  different 
caliber. 

1.  Provide  yourself  with  several  strong  threads  about  four  inches 
in  length  for  ligating  blood  vessels.     Have  the  red  injection  mass 
melted  and  heated  to  about  50°  C.     Also  have  ready  some  hot  water 
to  warm  the  syringe. 

2.  Kill  a  cat  or  a  rabbit  by  means  of  chloroform  or  illuminating 
gas.     The  latter  acts  more  rapidly  and  causes  less  struggle  on  the 
part  of  the  animal.     Work  rapidly  so  that  the  entire  animal  may  be 
injected  while  yet  warm.     Stretch  it  out  in  a  dissecting  pan  or  tie 
it  out  on  to  a  board,  or,  better,  keep  it  immersed  in  a  vessel  of  warm 
normal  saline  solution. 

3.  Slit  the  skin  along  the  ventral  surface  of  the  body  to  the  middle 
of  the  neck  and  reflect  it  to  the  right  and  left  side.     Pin  it  back  out 
of  the  way. 

4.  Snip  a  small  hole  through  the  body,  wall  just  posterior  to  the 
ensiform  cartilage.     Insert  the  index  finger  of  the  left  hand  to  guide 


Injection  of  Blood  and  Lymph  Vessels  85 

the  scissors  and  prevent  injury  to  the  underlying  organs,  and  cut 
the  costosternal  cartilages  of  the  right  side  up  to  the  first  rib.  In 
like  manner  cut  the  cartilages  of  the  left  side  up  to  the  first  rib. 

5.  Ligate  the  sternum  tightly  as  close  to  the  first  ribs  as  possible 
to  prevent  leakage  from  cut  blood  vessels. 

6.  Cut  off  the  apex  of  the  heart  and  expose  the  ventricles.     The 
left  ventricle  is  seen  as  a  round  opening,  the  right  as  a  slit. 

7.  With  a  sponge  wrung  out  of  warm  water  rapidly  absorb  the 
blood  from  the  thorax. 

8.  Choose  the  largest  cannula  that  the  aorta  will  admit  and  thrust 
it  through  the  left  ventricle  into  the  aorta. 

9.  With  a  pair  of  fine-pointed  forceps  (preferably  with  curved 
points)  pick  up  one  end  of  a  thread  for  ligating  and  carefully  work 
it  through  under  the  aorta  (do  not  mistake  the  vena  cava  superior 
for  the  aorta).     Tie  the  thread  around  the  aorta  over  the  cannula, 
making  a  double  or  surgeon's  knot.     Draw  it  tightly  on  the  cannula 
so  that  the  latter  will  be  held  firmly  in  place.     Run  another  thread 
through  under  the  aorta  and  have  it  in  readiness  to  ligate  the  aorta 
when  the  cannula  is  withdrawn. 

10.  Warm  the  syringe  by  sucking  hot  water  into  it  repeatedly, 
then  fill  it  and  the  cannula  with  the  warm  injecting  fluid. 

11.  Force  out  a  little  of  the  fluid  from  the  syringe  to  expel  all 
air,  and  connect  it  carefully  with  the  cannula. 

12.  Force  the  injecting  mass  into  the  blood  vessels  by  a  slow 
steady  pressure.     Begin  with  a  very  low  pressure,  so  that  the  large 
vessels  will  be  thoroughly  filled  before  the  mass  enters  the  capillaries. 
The  pressure  should  be  gradually  increased.     Avoid  sudden  increase 
of  pressure  or  too  strong  pressure,  for  either  may  cause  a  rupture 
of  the  blood  vessels  and  consequent  extravasation.     From  8  to  10 
minutes  is  about  the  time  required  to  make  a  good  injection  of 
the  cat. 

13.  Examine  the  intestines  and  the  gums  from  time  to  time  and 
also  the  inside  of  the  thigh  (from  which  the  skin  has-  been  reflected) ; 
they  should  be  deeply  colored  by  the  mass  before  the  injection  is 
complete.     If  the  mass  begins  early  to  flow  from  the  right  ventricle, 
the  ventricle  should  be  ligated.     In  any  event,  it  is  well  to  tie  the 


86  Animal  Micrology 

ventricle  a  few  minutes  before  completion  of  the  injection,  to  insure 
filling  of  all  blood  vessels. 

NOTE. — If  the  gums  remain  uncolored,  the  cannula  has  probably  been  forced 
past  the  arteries  which  lead  to  the  head.  In  such  a  case,  complete  the  injection 
of  the  trunk  and  then,  if  injected  tissue  from  the  head  region  is  desired,  cut 
obliquely  into  one  side  of  the  innominate  artery,  tie  a  cannula  in  place,  and 
inject  toward  the  head  as  in  the  case  of  the  aorta. 

14.  When  the  injection  is  complete,  shut  the  stop-cock,  ligate 
the  aorta,  or  clamp  it  with  pressure  forceps  beyond  the  end  of  the 
cannula  and  then  remove  the  latter. 

15.  Place  the  animal  in  cold  water  or  cold  alcohol  for  half  an 
hour,  then  remove  pieces  of  liver,  spleen,  pancreas,  stomach,  intes- 
tine, salivary  glands,  kidneys,  and  voluntary  muscles  and  harden 
in  strong  alcohol  or  in  10  per  cent  formalin. 

16.  When  sufficiently  hardened  transfer  the  objects  to  ether- 
alcohol  and  proceed  to  imbed  and  cut  in  ceiloidin  according  to  the 
method  already  given.     Make  longitudinal  sections  of  the  kidney 
parallel  to  its  flat  surface.     Cut  transverse  sections  of  liver,  stomach, 
and  intestine,  longitudinal  ones  of  the  muscle,  and  sections  passing 
longitudinally  through  the  hilum  of  the  salivary  glands  and  spleen. 
The  sections  should  not  be  under  30  microns  thick.     Mount  some 
unstained;    stain  others  in  diluted  Delafield's  hematoxylin  or  in 

hemalum. 

MEMORANDA 

1.  Apparatus  for  Continuous  Air-Pressure  Injections  is  now  provided  in 
many  laboratories.  If  a  regular  cylinder  for  air  pressure  is  not  present, 
however,  anyone  with  a  little  ingenuity  can  readily  fit  up  a  suitable  apparatus. 
A  carboy  or  large-mouthed  bottle  which  can  be  tightly  corked  will  answer 
as  a  chamber  for  compressed  air,  a  water  tap,  or  a  tank  of  water  elevated  to 
the  height  of  7  or  8  feet  will  provide  sufficient  pressure.  By  making  the 
proper  connections  by  means  of  rubber  and  glass  tubing  a  steady  stream  of 
compressed  air  may  finally  be  conducted  to  a  flask  containing  the  injection 
mass;  the  flask  works  in  the  same  way  as  an  ordinary  wash-bottle  (Fig.  35). 
All  corks  and  fittings  must  be  tightly  secured  with  wire  or  strong  cord.  If 
desired,  by  adding  an  extra  perforation  to  the  cork  in  the  air  chamber,  a 
mercury  manometer  may  be  added  to  register  the  amount  of  air  pressure.  If 
a  metal  cannula  is  not  at  hand  a  glass  one  may  be  made  as  indicated  under 
memorandum  9,  p.  90.  In  lieu  of  a  stop-cock,  use  a  pinch-cock  on  the  rubber 
delivery  tube. 


Injection  of  Blood  and  Lymph  Vessels 


87 


2.  A  Double  Injection  of  the  Vascular  System  may  be  made  by  first 
injecting  the  blue  mass  until  it  is  seen  to  flow  from  the  right  ventricle,  then 
detaching  the  tube  which  conveys  the  blue  mass,  and  slipping  over  the  end 
of  the  cannula  a  tube  conveying  a  red  mass.  This  second  mass  should  be 
in  a  bottle  or  flask  connected  with  the  pressure  bottle  by  means  of  an  addi- 
tional tube  through  the  cork  of  the  latter,  or  the  two-  flasks  containing  the 
colored  masses  may  be  connected  with  the  tube  from  the  pressure  bottle  by 
means  of  a  Y-tube.  Each  must  be  provided  with  a  pinch-cock  or  clamp 
to  hold  back  its  contents  while  the  other  is  in  operation.  If  a  syringe  is 
used,  it  is  better  to  have  a  second  syringe  for  the  second  mass,  although  one 
will  answer  if  it  is  rinsed  out  with  hot  water  before  being  filled  with  the  sec- 
ond mass.  The  second  mass  should  have  a  quantity  of  very  finely  pulverized 


PIG.  35. — Apparatus  for  Continuous  Air-Pressure  Injection  (after  B.  G.  Smith) 

cornstarch  mixed  with  it,  so  that  when  it  reaches  the  capillaries  they  will 
become  completely  plugged.  Walker  uses  the  red  gelatin  mass  first,  then 
follows  with  a  gelatin  colored  with  ultramarine  blue.  The  granules  of  the 
atter  are  too  large  to  enter  the  capillaries,  hence  a  double  injection  with 
veins  red  and  arteries  blue  is  obtained. 

It  should  be  borne  in  mind  that  the  larger  veins  cannot  be  injected  in  a 
direction  contrary  to  their  flow  because  of  the  valves  they  contain. 

3.  The  Lungs,  Liver,  and  Kidneys  are  readily  injected  through  then- 
larger  blood  vessels  with  two  masses,  and  afford  very  instructive  material 
when  thus  prepared.  A  triple  injection  of  the  liver  may  be  made  by  inject- 
ing the  hepatic  artery  and  the  hepatic  and  portal  veins.  The  third  mass 
may  be  colored  with  China  ink.  Whitman  (Methods  in  Microscopical 
Anatomy  and  Embryology)  recommends  first  injecting  the  hepatic  artery 


88  .  Animal  Micrology 

and  afterward  the  two  veins.     The  blood  should  be  washed  out  of  the  organ 
to  be  injected,  with  warm  salt  solution. 

4.  To  Inject  Lymphatics,  the  puncture  method  is  commonly  employed. 
For  example,  an  aqueous  solution  of  Berlin  blue  is  drawn  into  a  hypodermic 
syringe,  the  sharp  point  of  the  cannula  is  thrust  into  the  tissue,  and  the 
syringe  emptied  by  slight,  steady  pressure.     For  practice,  thrust  the  cannula 
into  the  pad  of  a  cat's  foot,  and  force  in  some  of  the  injection  mass.     If  the 
leg  is  rubbed  upward,  the  fluid  will  flow  along  the  lymph  channels  and  into 
the  glands  of  the  groin.     Instead  of  this  haphazard  method,  however,  much 
better  results  will  be  insured  by  the  use  of  the  needle-and-clamp  device  of 
W.  S.  Miller  (Johns  Hopkins  Hospital  Bulletin,  XVI,  No.  173  [1905]). 

5.  Micro-Injection  of  Embryonic  Vessels  has  been  much  resorted  to  in 
recent  years  for  the  study  of  early  stages  of  lymph  and  blood  vessels,  and 
some  very  delicate  and  effective  methods  have  been  devised.     India  ink  is 
the  medium  ordinarily  used.    Embryos  of  medium  size  are  generally  injected 
with  a  hypodermic  syringe,  but  smaller  embryos  require  a  more  delicate 
procedure  in  which  glass  tubes  with  the  finest  possible  capillary  points  are 
used. 

Anyone  who  has  seen  the  beautifully  injected  specimens  of  Dr.  H.  McE. 
Knower  will  concede  the  success  of  his  method.  The  general  scheme  of 
his  apparatus  is  shown  in  Fig.  36,  and  the  accompanying  legend  is  self- 
explanatory. 

He  expresses  the  essence  of  his  method  as  follows:  "If  a  gentle  warmth 
is  applied  to  a  glass  bulb  blown  on  the  end  of  a  capillary  tube,  while  the  fine 
point  of  the  tube  is  held  beneath  the  surface  of  some  fluid,  such  as  India 
ink,  air  will  be  driven  out  of  the  bulb  and  ink  will  run  up  to  replace  it  as  the 
bulb  cools.  When  the  system  has  come  to  equilibrium,  the  point  of  the  tube 
is  inserted  into  the  desired  blood  vessel  under  a  dissecting  microscope 
(binocular  if  possible),  the  tube  being  carried  on  a  holder  to  avoid  warming 
it  or  the  bulb.  India  ink  is  now  injected,  as  desired,  by  warming  the  bulb 
when  ready."  The  method  is  explained  in  minute  detail  in  the  Anotomical 
Record  for  August,  1908  (Vol.  II,  No.  5),  together  with  applications  to  fish, 
amphibia,  reptiles,  birds,  and  mammals. 

Instead  of  using  a  glass  bulb,  as  does  Knower,  Heuser  inserts  a  simply 
made  resistance  coil  of  fine  German-silver  wire  into  an  ordinary  salt-mouth 
bottle.  The  ends  of  the  wire  extend  out  through  the  cork  for  electrical 
connection.  When  a  current  is  passed  through  the  wire  the  air  surrounding 
it  in  the  bottle,  becoming  heated,  expands  and  affords  a  steady  and  pro- 
longed pressure. 

Dr.  Emily  Ray  Gregory  uses  direct  pressure  from  a  good  grade  of 
De  Vilbiss'  atomizer  bulb  which  lies  on  the  floor.  The  bulb,  kept  from 
rolling  by  a  crocheted  net  cover,  is  operated  by  the  foot.  Pressure  is  trans- 
mitted through  a  i^-inch  red-rubber  tube  to  the  short  glass  injection  tube 


Injection  of  Blood  and  Lymph  Vessels 


89 


FIG.  36. — Apparatus  for  the  Injection  of  Small  Embryos,  etc.,  under  the  Microscope 
(after  Knower). 

1.  Glass  tube  drawn  out  after  moderate  heating  near  middle  (see  arrow).     Tube 
should  be  turned  while  heating.     The  capillary  tube  will  usually  be  three  or  four  times 
the  length  of  that  in  the  figure. 

2.  Superfluous  tube  burned  off  at  x  by  a  fine  blow-pipe  flame.     Tube  should  be 
turned  while  heating. 

3.  Enlarged  end,  molten  and  heated  around  the  end  (note  arrow)  while  being  turned. 

4.  Bulb  blown  as  soon  as  tube  in  Fig.  3  is  molten.    The  end  is  usually  removed  from 
the  flame,  when  hot  enough,  and  blown  quickly,  but  without  too  great  pressure,  through 
the  capillary  tube.     Size  usually  about  twice  that  of  figure. 

5.  The  bend  at  a  is  given  before  the  fine  tip  b  is  drawn  out  over  the  gentle  heat  of  c, 
or  near  a  hot  iron.     Reduced  size. 

6.  Board  with  auger  holes  to  hold  bulbs.     Size  much  reduced. 

7.  Method  of  applying  heat  and  introducing  tip  with  free  hand,  and  simple  cork 
holder.     Size  reduced. 

8.  Wire  support  w,  for  holding  bulb  while  filling.     Reduced  size. 

9.  Method  of  using  jointed  holder  a,  with  binocular.     The  small  gas-jet  c  lies  in 
front  ready  for  heating  the  bulb.     Size  greatly  reduced. 


90  Animal  Micrology 

which  has  the  outer  end  drawn  into  a  capillary  tip  at  right  angles  to  itself. 
The  method  is  given  in  detail  in  the  August  number  of  the  Anatomical 
Record,  1916  (Vol.  XI,  No.  1). 

For  other  suggestions  regarding  micro-injections  see  Hoyer,  Zeitschrift 
fur  wissenschaftliche  Mikroskopie,  Band  XXV  (1908);  Evans,  American 
Journal  of  Anatomy,  IX  (1910);  and  Sabin,  Contributions  to  Embryology, 
III,  No.  7  (1915),  Carnegie  Institution  of  Washington. 

6.  To  Keep  Gelatin  Injection  Masses  let  them  congeal,  then  cover  the 
surface  with  95  per  cent  alcohol,  and  leave  in  a  well-stoppered  vessel  until 
needed. 

7.  Injection  through  the  Femqral  Artery  is  frequently  practiced,  and 
is  preferred  to  injection  through  the  aorta  by  some  workers.    An  oblique 
cut  is  made  in  one  side  of  the  artery  and  the  cannula  inserted  pointing 
toward  the  heart.    Others  prefer  to  cut  into  the  dorsal  aorta  and  inject 
both  anteriorly  and  posteriorly. 

8.  The  Injecting  Syringe  must  work  without  jerking  or  catching  along  the 
wall  of  the  barrel.     It  should  always  be  carefully  cleaned  after  using.    If  the 
piston  does  not  fit  the  barrel  tightly  enough  it  should  be  wrapped  with  gauze. 

9.  Glass  Cannulae  may  be  made  by  grasping  the  ends  of  a  short  piece 
of  soft  glass  tubing  and  heating  the  middle  in  a  flame  until  the  glass  becomes 
soft,  which  is  indicated  by  the  yellow  color  of  the  flame.    The  tubing  should 
be  constantly  rotated,  so  that  all  sides  heat  equally.    When  the  glass  becomes 
soft,  draw  the  tube  out  steadily  until  the  diameter  of  the  soft  portion 
becomes  as  small  as  desired.    When  the  glass  has  cooled,  the  tube  should 
be  cut  with  a  file  at  the  proper  place  to  make  two  cannulae  of  it. 

10.  If  the  Blue  Color  Fades  in  the  gelatin  mass  in  the  tissues,  it  may 
frequently  be  restored  by  treating  the  tissue  or  section  with  oil  of  cloves  or 
turpentine. 

11.  A  Cold  Fluid  Gelatin  Mass  has  been  used  successfully  by  Tandler  (see 
abstract  by  A.  M.  C.  in  Journal  of  Applied  Microscopy,  V,  1625).     To  pre- 
pare the  mass,  dissolve  5  grams  of  finest  gelatin  in  100  c.c.  of  tepid  distilled 
water.     Color  to  the  desired  shade  with  Berlin  blue,  and  then  add  slowly 
5  to  6  grams  of  potassium  iodide.     The  mass  remains  fluid  at  ordinary 
temperatures,  but  when  injected  objects  are  placed  in  5  per  cent  formalin 
it  sets  completely  and  is  thereafter  unaffected  by  reagents.    The  minutest 
vessels  are  injected,  and  sections  may  be  stained  in  the  usual  ways.    Sub- 
jection to  strong  acids,  such  as  sulphuric  or  hydrochloric,  does  not  affect 
the  mass;  hence  it  may  be  used  for  injecting  specimens  that  are  to  be  decal- 
cified afterward.     To  preserve  the  fresh  mass,  add  a  few  crystals  of  thymol 
and  keep  in  a  stoppered  bottle. 

12.  Corrosion  of  Injected  Vessels  or  Cavities  is  sometimes  practiced. 
A  mass  must  be  employed  which  will  not  be  attacked  by  the  reagent  used 


Injection  of  Blood  and  Lymph  Vessels  91 

for  destroying  the  surrounding  tissues.  One  of  the  best  masses  consists  of 
white  wax  5  parts  and  rosin  6  parts,  melted  together  at  a  tempera- 
ture of  about  75°  C.  For  fine  vessels  increase  the  proportions  of  wax,  for 
larger  ones  add  more  rosin.  Vermilion,  Prussian  blue,  or  "chromate 
of  lead  may  be  used  for  coloring.  The  part  to  be  injected  should  be  placed 
in  warm  water  and  the  mass  injected  at  a  temperature  of  from  50°  to  60°  C. 
The  injected  part  is  left  in  cold  water  for  from  1  to  2  hours,  and  is  then  cor- 
roded in  pure  hydrochloric  acid  for  from  6  to  48  hours,  according  to  the 
resistance  of  the  tissue.  Finally,  wash  the  preparation  thoroughly  in 
running  water.  For  bibliography  and  more  detailed  directions  see 
Technique  des  injections,  by  Hermann  Joris,  Universite  Libre  de  Bruxelles, 
1903. 

13.  Wood's  Metal  is  one  of  the  commonest  injection  media  used  for 
corrosion  preparations.     It  is  a  fusible  alloy  consisting  of  1  cr  2  parts  of 
cadmium,  2  parts  of  tin,  4  of  lead,  with  7  or  8  parts  of  bismuth.     It  melts 
at  from  66°  to  71°  C. 

14.  Celluloid  Dissolved  in  Acetone  was  found  preferable  by  Flint 
(American  Journal  of  Anatomy,  VI  [1906]),  in  his  study  of  lung  develop- 
ment, to  celloidin  or  Wood's  metal  for  corrosion  preparations.    He  injected 
from  aspkation  bottles  into  the  lungs  through  the  trachea,  and  used  hydro- 
chloric acid  for  corrosion. 

15.  Air  Injection  of  Minute  Vessels  was  found  to  be  an  indispensable 
method  by  Locy  (American  Journal  of  Anatomy,  XIX  [May,  1916],  3)  in 
his  work  on  the  lung  and  air  passages  of  the  chick:   "In  stages  subsequent 
to  96  hours,  the  lungs  and  air  sacs  were  dissected  out  of  the  previously 
fixed  and  hardened  specimens,  then  cleared  in  cedar  oil,  after  which  the 
organs  were  placed  in  a  mixture  of  1  part  cedar  oil  and  2  parts  chloroform. 
On  becoming  permeated  with  this  fluid,  the  preparation  was  removed  from 
the  mixture  and  placed  on  a  filter  paper  until  the  chloroform  might  evaporate. 
The  evaporation  of  the  chloroform  served  to  draw  the  cedar  oil  from  the 
lumina  of  the  various  branches  of  the  bronchial  tree  into  the  lung  tissue  and 
to  fill  the  spaces  thus  made  with  air.     When  this  preparation  was  replaced 
in  pure  cedar  oil,  the  difference  between  the  refractive  index  of  the  im- 
prisoned air  and  the  surrounding  medium  gave  the  lung  tubes  the  appearance 
of  being  filled  with  a  metallic  cast.    Thus  the  minute  air  passages  that 
could  not  be  injected  by  other  means  were  made  clear.    The  finer  details 
would   disappear  after  a  few  minutes  as  the  cedar  oil  percolated  into 
them,  but  the  same  specimen,  if  carefully  manipulated,  can  be  treated 
repeatedly  without  apparent  injury,  and  a  complete  picture  could  finally  be 
obtained." 

For  later  stages  Locy  also  used  celloidin  and  Wood's  metal  injections 
followed  by  corrosions.    He  has  obtained  some  beautiful  Wood's  metal 


92  Animal  Micrology 

casts  of  the  adult  lung.  The  lungs  of  the  freshly  killed  fowl  were  distended 
under  pressure  with  80  per  cent  alcohol  until  the  air  sacs  were  fully  expanded. 
The  entire  bird  was  then  immersed  in  alcohol  for  24  hours  or  more  before 
metallic  injection  was  attempted. 

16.  An  Excellent  Injection  Mass  for  other  than  histological  purposes  as 
used  in  our  own  laboratories  is  made  as  follows  (Wagner) : 

Water 100  c.c. 

Glycerin 20  c.c. 

Strong  formalin 20  c.c. 

Cornstarch,  powdered 75  grams 

Mix  by  gradually  adding  water  and  glycerin  to  the  starch,  rubbing  out 
all  lumps.  For  yellow  color  add  10  grams  of  chrome  yellow;  for  green, 
10  grams  of  chrome  green;  for  red,  10  grams  of  vermilion.  Strain  through 
cheesecloth  and  add  the  formalin.  If  the  mass  is  too  thick  to  strain,  add  the 
formalin  first.  Cornstarch  is  vastly  superior  to  laundry  starch  and  the 
colors  recommended  diffuse  less  into  tissues  than  carmine  or  Berlin  blue. 

17.  For  Clearing  of  Injected  Organs  or  Embryos  "in  Toto,"  see  mem- 
oranda 15,  16,  and  17  (pp.  102-104). 


CHAPTER  XIII 

OBJECTS  OF  GENERAL  INTEREST:   CELL-MAKING,  FLUID 
MOUNTS,  "IN  TOTO"  PREPARATIONS,  DRY  MOUNTS, 
OPAQUE  MOUNTS 

When  objects  of  considerable  thickness  are  to  be  mounted,  it 
is  sometimes  necessary  to  resort  to  cells  which  will  contain  the  object 
and  support  the  cover-glass.  Fluid  mounts  and  aqueous  media 
must  occasionally  be  used  for  delicate  objects  which  would  be  injuri- 
ously affected  by  alcohol,  or  which  are  unsuitable  for  mounting  in 
balsam.  When  such  mounts  are  used,  whether  in  a  cell  or  not,  the 
cover-glass  must  ordinarily  be  sealed  with  a  cement  if  the  prepara- 
tion is  to  be  permanent.  In  all  cases  where  it  is  at  all  practicable, 
balsam  mounts  are  to  be  preferred  for  permanent  preparations. 
Glycerin  is  a  convenient  mounting-medium  for  many  objects,  espe- 
cially for  temporary  mounts.  It  is  often  used  where  such  media 
as  balsam  would  render  the  preparation  too  transparent;  it  is  much 
more  favorable,  moreover,  to  the  preservation  of  color  than  are 
resinous  media.  For  making  cells  and  sealing  circular  covers,  a 
turntable  (Fig.  37)  is  desirable,  although  the  work  may  be  done  by 
following  a  guide  ring  drawn  on  paper  and  placed  under  the  slide. 

I.    TURNING  CELLS 

Prepare  12  or  15  slides  as  follows:  1.  Place  a  slide  on  a  turn- 
table and  adjust  it  so  that  its  center  lies  over  the  center  of  the  turn- 
table. 

2.  Dip  a  small  camel's  hair  pencil  into  gold  size,  but  do  not 
take  up  enough  of  the  fluid  to  drop  (see  also  memorandum  13, 
p.  101). 

3.  Choose  a  guide  ring  on  the  turntable  which  is  of  slightly 
smaller  diameter  than  the  cover-glass  to  be  used,  whirl  the  table 
and  hold  the  pencil  lightly  over  the  guide  ring.     The  ring  which 

93 


94  Animal  Micrology 

has  been  spun  should  be  even.  If  it  is  not,  practice  turning  rings 
until  satisfactory  ones  are  made.  If  the  gold  size  is  old  it  is  prob- 
ably too  thick  to  make  suitable  rings.  Pure  linseed  oil  may  be  used 
to  dilute  it,  but  it  is  advisable  to  use  only  fresh  gold  size  if  it  is 
obtainable. 

4.  The  slide  must  be  set  aside  to  dry  before  it  can  be  used  for 
mounting.     A  gentle  heat  will  aid  in  drying. 


PIG.  37. — Turntable 

5.  To  some  of  the  cells  add  successive  coats  of  gold  size  as  the 
previous  one  dries,  so  that  you  will  have  cells  of  varying  depth. 

II.    MOUNTING  IN  GLYCERIN 

A.  Water  Mites  and  Transparent  Larvae.— 1.  Kill  several  small, 
colored  water  mites  or  transparent  larvae  of  insects  by  means  of 
chloroform  (a  few  drops  in  water)  and  place  them  for  half  an 
hour  (two  or  three  hours  for  larger  objects)  into  a  mixture  of 
water  and  glycerin  equal  parts,  after  which  transfer  them  to  pure 
glycerin. 

2.  Apply  a  thin  coat  of  gold  size  to  the  upper  edge  of  a  cell  which 
is  of  sufficient  depth  to  accommodate  the  object. 

3.  Breathe  into  the  cell  to  moisten  it  so  that  the  glycerin  will 
adhere  throughout  and  prevent  the  formation  of  air  bubbles. 


Objects  of  General  Interest  95 

4.  Fill  the  cell  flush  with  glycerin  and  put  the  object  into  it,  care- 
fully spreading  out  all  parts. 

5.  Breathe  on  the  lower  surface  of  a  clean  cover-glass,  put  one 
edge  down  on  the  edge  of  the  cell,  and  then  gradually  lower  the  cover 
so  as  to  avoid  bubbles  of  air.     When  in  place,  press  the  cover  down 
gently  with  the  handle  of  a  needle  and  see  that  it  adheres  all  around. 
Wash  off  the  exuded  glycerin  and  carefully  wipe  the  slide  with  a 
cloth. 

6.  Turn  a  comparatively  broad  ring  of  gold  size  around  the 
edge  of  the  cover  to  seal  it,  and  when  this  is  dry  add  a  very  thin 
coat  of  Bell's  cement.     Label  and  put  away  in  a  horizontal  position 
until  dry. 

CAUTION. — It  is  indispensable  that  the  edges  of  the  cover-glass 
be  perfectly  dry  before  attempting  to  seal  the  preparation;  other- 
wise the  cement  will  not  adhere. 

B.  Killing  and  Mounting  Hydra. — 1.  With  a  dipping-tube 
(memorandum  10,  p.  101)  remove  a  hydra  to  a  warm  watch-glass  and 
leave  it  in  only  a  few  drops  of  water.  Have  ready  some  hot  Bouin's 
fluid  or  corrosive  acetic,  and  when  the  hydra  sends  out  its  tentacles 
and  expands  its  body,  apply  the  reagent  by  suddenly  squirting  it  into 
the  watch-glass  so  that  it  sweeps  over  the  hydra  from  aboral  to 
oral  extremity  and  carries  the  tentacles  out  straight.  Then  fill  the 
watch-glass  with  the  hot  fluid. 

2.  After  10  minutes  pour  off  the  fixing  fluid  and  .wash  the  animal 
thoroughly  in  70  per  cent  alcohol. 

3.  Replace  the  alcohol  with  alum-cochineal  or  dilute  hema- 
toxylin  and  stain  for  from  30  minutes  to  several  hours. 

4.  Remove  the   stain   with  a   pipette  and  replace  it  with  a 
rnixture  of  equal  parts  of  glycerin  and  water  for  half  an  hour, 
followed  by   pure  glycerin.     Proceed  farther  as  in  the  preceding 
exercise. 

NOTE. — After  removal  from  the  stain,  if  necessary,  decolorize  in  acidulated 
water  or  alcohol  (0.5  per  cent  hydrochloric  acid),  then  wash  out  the  acid  thor- 
oughly in  tap  water. 

Hydra  may  also  be  dehydrated,  cleared,  and  mounted  in  balsam 
(see  also  "Hydra,"  p.  264). 


96  Animal  Micrology 

HI.    MOUNTING  IN  GLYCERIN- JELLY 

Glycerin-jelly  is  frequently  preferable  to  pure  glycerin  for  mount- 
ing because  it  is  a  solid  at  ordinary  temperatures.  One  formula  for 
making  it  is  as  follows : 

Water 42  c.c. 

Gelatin 6  grams 

Glycerin 50  c.c. 

Carbolic-acid  crystals 2  grams 

Let  the  gelatin  soak  in  the  water  for  half  an  hour,  then  dissolve 
with  gentle  heat.  Add  about  5  c.c.  of  white  of  egg  and  heat  (not 
over  75°  C.)  for  half  an  hour.  The  egg  albumen  gradually  precipi- 
tates and  carries  down  all  fine  particles  of  dust,  etc.,  so  that  the 
gelatin  is  left  perfectly  clear. 

Filter  through  moist,  fine  hot  flannel  and  add  the  glycerin  and  the 
carbolic  acid.  Use  only  clean  gelatin  of  the  best  quality.  Warm 
for  10  or  15  minutes,  stirring  continually  until  the  mixture  is 
homogeneous.  If  heated  above  75°  C.,  the  gelatin  may  be  trans- 
formed into  metagelatin,  which  will  not  harden  at  ordinary 
temperatures. 

A.  Small  Crustacea. — 1.  By  means  of  a  dipping-tube  isolate  such 
small  creatures  as  Cyclops,  Daphnia,  or  Cypris. 

2.  Kill  by  warming  slowly  in  a  drop  of  water  on  a  slide. 

3.  Place  them  in  a  cell  of  proper  depth,  draw  off  all  water  with  a 
pipette,  and  gently  warm  the  slide. 

4.  Place  the  bottle  of  glycerin-jelly  into  a  vessel  containing  warm 
water  until  the  jelly  becomes  liquid,  but  do  not  let  it  get  any  warmer. 

5.  Fill  the  cell  flush  with  the  warm  jelly  and  arrange  the  objects  in 
suitable  positions. 

6.  Breathe  upon  the  lower  surface  of  a  clean  cover-glass  and 
put  it  in  place  in  the  usual  way. 

7.  Wash  away  any  trace  of  the  jelly  from  the  outside  of  the  cell 
and  when  the  slide  is  dry  run  a  ring  of  gold-size  cement  around  the 
edge  of  the  cover.     After  this  dries,  varnish  with  Bell's  cement.     It 
is  not  an  absolute  necessity  to  seal  glycerin-jelly  mounts,  but  the 
writer  has  always  found  it  a  wise  precaution. 


Objects  of  General  Interest  97 

B.  Muscle  of  Insect. — 1.  Cut  off  the  head  of  an  insect  and  bisect  the 
trunk  so  as  to  expose  the  interior.  Observe  two  kinds  of  muscular  tissue, 
that  of  grayish  color  belonging  to  the  legs,  the  yellowish  to  the  wings. 

2.  Take  a  shred  of  muscle  and  on  a  dry  slide  carefully  separate  pieces 
of  muscle  fiber  and  stretch  them  out,  while  keeping  them  moist  by  breathing 
on  them. 

3.  Mount  in  glycerin-jelly  as  directed  in  the  previous  exercise  (see  also 
p.  251). 

IV.    MOUNTING  IN  BALSAM 

A.  Flat  Worms. — 1.  Obtain  specimens  of  Planaria  from  the 
under  surface  of  flat  rocks  in  the  edge  of  streams  (see  "Planaria," 
p.  266). 

2.  Place  the  animal  in  a  little  tepid  water.     Watch  until  it  is 
extended  full  length,  then  flood  it  quickly  with  corrosive  sublimate 
to  which  1  to  3  per  cent  of  acetic  acid  has  been  added.     The  animal 
may  be  removed  after  30  minutes  or  an  hour  and  washed  thoroughly  in 
50  per  cent  alcohol  to  which  a  little  tincture  of  iodine  has  been  added. 

3.  Stain  for  24  hours  in  alum-cochineal  or  in  Delafield's  hema- 
toxylin  diluted  one-half  with  water. 

4.  Wash  in  water  followed  by  35  and  50  per  cent  alcohol  each  15 
minutes. 

5.  Decolorize  in  acid  alcohol  until  the  color  ceases  to  come  away 
freely  (10  to  30  minutes). 

6.  Wash  out  the  acid  in  70  per  cent  alcohol,  using  the  alkaline 
alcohol  if  hematoxylin  was  used  in  staining. 

7.  Flatten  the  animal  by  compressing  it  between  two  slides  by 
means  of  a  rubber  band,  and  place  it  for  24  hours  in  95  per  cent 
alcohol. 

8.  Transfer  to  absolute  alcohol  for  1  hour,  and  to  xylol  until  clear. 

9.  Mount  in  balsam  in  a  thin  cell  or  without  a  cell  at  pleasure. 
If  on  examination  the  separate  organs  of  the  animal  are  not  seen 
distinctly,  it  probably  has  not  been  compressed  sufficiently.     This 
difficulty  may  sometimes  be  avoided  in  a  measure  by  letting  a 
cover-glass  rest  upon  the  live  planarian  to  flatten  it  out  slightly,  and 
then  running  the  fixing  fluid  under  the  cover.     Specimens  which 
have  been  in  the  laboratory  for  some  weeks  or  months  make  better 
preparations  than  those  fresh  from  the  stream. 


98  Animal  Micrology 

B.  Mosquito,  Gnat,  or  Aphid. — 1.  Kill  a  mosquito  with  cyanide 
or  chloroform  and  place  it  in  cedar  oil  or  turpentine  for  an  hour. 

2.  Remove,  and  place  it  on  its  back  on  filter-paper.    Carefully 
spread  the  legs  of  the  insect,  put  a  drop  of  thick  balsam  on  a  slide, 
invert  the  slide,  and  bring  the  balsam  in  contact  with  the  thorax  of  the 
mosquito.     Spread  the  wings  and  the  legs  of  the  insect  and  gently 
press  it  down  into  the  balsam. 

3.  Add  thinner  balsam,  see  that  the  proboscis  and  antennae 
are  floated  out  properly,  then  add  more  balsam,  and  put  on  a  cover- 
glass. 

V.    OPAQUE  MOUNTS 

Some  objects  are  mounted  to  be  viewed  by  reflected  instead  of  trans- 
mitted light.  They  may  be  mounted  in  the  ordinary  way,  and  when  they 
are  examined  as  opaque  objects,  the  light  from  the  mirror  should  be  turned 
away  and,  if  necessary,  a  strip  of  dark  paper  placed  under  the  slide  to  shut 
off  all  light  from  below. 

A.  Beetles. — Choose  a  shallow  cell  for  mounting  the  wing  cases  and  legs 
of  one  of  the  Curculionidae,  preferably  Curculio  imperalis,  the  South  Ameri- 
can diamond  beetle. 

1.  Soak  the  part  in  cedar  oil  or  turpentine  for  half  an  hour,  then  place  it  in 
the  cell  in  the  proper  position,  the  outer  side  of  the  case  toward  the  observer. 

2.  Fill  up  the  cell  with  balsam  and  add  the  cover. 

B.  Wings  of  Moths  or  Butterflies. — Prepare  parts  of  the  wings  of  moths 
or  butterflies  as  in  A.    The  wing  of  the  clothes  moth  makes  a  good  opaque 
mount. 

C.  Head  of  a  Fly. — 1.  Secure  the  specimen  (preferably  one  having 
colored  eyes,  as  one  of  the  gadflies)  and  choose  a  cell  of  the  proper  size  for 
it.    The  cell  should  be  of  such  a  depth  that  the  cover  will  rest  lightly  upon 
the  object  and  retain  it  in  the  center  of  the  cell.    The  head  should  present 
the  front  view  when  mounted. 

2.  Spin  a  very  thin  coat  of  gold  size  on  to  the  dry  edge  of  the  cell  so  that 
the  cover  will  adhere. 

3.  Soak  the  head  of  the  fly  for  a  couple  of  hours  in  equal  parts  of  glycerin 
and  water. 

4.  Moisten  the  cell  by  breathing  into  it,  fill  it  with  glycerin,  and  transfer 
the  object  to  it. 

5.  Breathe  on  the  cover-glass  and  apply  it  very  carefully  to  avoid  air 
bubbles.    When  the  cover  settles  into  place,  press  it  down  gently  to  make 
it  adhere  to  the  cement. 

6.  Set  it  aside  to  harden.    When  hard,  seal  on  the  turntable  with  gold 
size  followed  by  Bell's  cement  when  the  gold  size  is  dry. 


Objects  of  General  Interest  99 

D.  Foreleg  of  Dytiscus,  the  Great  Water  Beetle. — 1.  Detach  the  foreleg 
of  a  male,  and  soak  it  in  10  per  cent  potash  solution  (see  reagent  86,  p.  237) 
for  a  day  or  two. 

2.  Wash  it  in  water,  run  it  up  to  95  per  cent  alcohol,  and  leave  it  there 
for  24  hours. 

3.  Pass  it  through  absolute  alcohol  and  clear  in  cedar  oil,  turpentine, 
or  xylol. 

4.  Lay  the  leg,  disk  side  uppermost,  in  a  drop  of  balsam  on  a  slide,  add 
another  drop  of  balsam,  and  carefully  cover  with  a  clean  cover-glass.    Place 
a  small  weight  (e.g.,  half  of  a  bullet)  on  top  of  the  cover  to  hold  it  down  until 
the  balsam  hardens. 

VI.    DRY  MOUNTS 

A.  Scales. — Prepare  a  very  shallow  cell  and  let  it  dry.    Thoroughly 
dry  the  scales  from  a  moth's  wing  by  gently  heating  them  on  a  slide  over  a 
flame.    Place  the  scales  in  a  cell,  warm  the  slide  until  the  cell  wall  becomes 
sticky,  put  on  the  cover  and  press  it  down  until  it  adheres  all  around,  and 
finally  seal  as  in  previous  exercises. 

B.  Eggs  of  Butterflies,  Small  Feathers,  Antennae  of  Insects,  etc.,  may 
be  mounted  as  dry  objects.    Care  must  be  taken  to  have  them  perfectly 
dry,  or  they  will  in  time  cloud  the  cover  with  moisture  from  within. 

MEMORANDA 

1.  Small  or  Soft  Insects  or  Their  Larvae  may  frequently  be  mounted 
directly  in  glycerin,  or  they  may  be  dehydrated  and  mounted  in  balsam. 
A  method  often  used  is  to  kill  them  in  strong  carbolic  acid  and  mount  them 
directly  in  balsam.    The  carbolic  acid  both  dehydrates  and  clears.    It  is 
better,  however,  to  clear  the  preparation  further  by  immersion  in  cedar  oil 
or  xylol  before  adding  the  balsam. 

2.  Insects  Having  Hard  Shells  must  first  be  soaked  in  10  per  cent  potash 
to  soften  them  and  render  them  transparent  if  they  are  to  be  examined 
by  transmitted  light.    The  softer  parts  of  insects  so  treated  are  destroyed 
and  only  the  external  parts  remain.    Such  insects  may  be  mounted  in  glycerin 
or  glycerin- jelly,  or  they  may  be  dehydrated,  cleared,  and  mounted  in 
balsam. 

3.  Delicate  Insects  which  are  too  frail  to  withstand  much  handling 
may  be  placed  at  once  in  cedar  oil  or  turpentine  and  after  an  hour  mounted 
in  balsam  (see  "Mosquito,"  p.  98). 

4.  Wings,  Legs,  Antennae,  Mouth-Parts,  etc.,  of  Such  Forms  as  Flies 
and  Bees,  which  have  been  preserved  in  alcohol,  should  be  completely 
dehydrated,  cleared,  and  mounted  in  balsam  in  cells  of  the  proper  depth. 

5.  Transparent  and  Soft  Insects  may  be  stained  in  alum-cochineal  or 
heinatoxylin  in  the  ordinary  way  and  mounted  as  whole  objects,  if  desired. 
They  will  stain  better  if  they  have  been  fixed  previously  in  some  corrosive- 


100  Animal  Micrology 

sublimate  mixture  and  then  washed  properly  (see  reagent  14,  p.  212).    To 
stain,  follow  the  method  outlined  in  IV,  A,  p.  97. 

6.  To  Center  an  Object  in  a  Cell  (the  head  of  an  insect,  for  example), 
thread  a  fine  needle  with  a  hair  and  run  it  through  the  object.     Remove  the 
needle  and  imbed  the  ends  of  the  hair  in  the  cement  on  opposite  sides  of  the 
cell.    When  the  cover-glass  is  put  in  place  the  object  may  be  adjusted  by 
pulling  the  hair.    After  the  slide  is  finished  and  dry,  the  ends  of  the  hair 
should  be  cut  off  at  the  edge  of  the  cell. 

Another  method  which  will  frequently  answer  for  an  object  to  be  mounted 
in  balsam  is  to  place  the  object  (after  clearing)  in  the  center  of  the  cell, 
coat  it  with  balsam,  adjust  it  properly,  and  then  set  the  slide  away  in  a 
place  free  from  dust  till  the  balsam  thickens.  Finally  fill  the  cell  with 
balsam  and  add  the  cover. 

7.  The  Radula  or  Lingual  Ribbon  of  the  Snail  or  Slug  should  be  dissected 
out  and  soaked  for  a  day  or  two  in  a  10  per  cent  solution  of  potash.    If  the 
animal  is  a  small  one,  cut  off  the  head  including  the  buccal  mass  and  soak  it 
in  a  solution  of  potash  until  the  soft  tissues  are  destroyed  and  only  the 
radula  remains.     From  the  potash  the  radula  is  transferred  to  water  and 
washed  for  some  hours.    With  a  strip  of  paper  on  each  side  to  prevent 
crushing  it,  it  should  be  placed  between  two  slides,  and  the  slides  bound 
together  by  means  of  string  or  rubber  bands.    While  held  in  this  position, 
dehydrate  and  clear  it.    Finally  remove  one  slide  and  the  paper  and  mount 
the  object  in  balsam  on  the  other  slide.    A  shallow  cell  may  be  used  if 
desired. 

8.  Flukes  and  Tapeworms  are  prepared  in  the  same  manner  as  Planaria 
(p.  97).    The  time  of  immersion  in  the  various  fluids  should  be  lengthened 
in  proportion  as  the  object  is  larger  than  the  planarian.     See  also  p.  267. 

For  in  toto  staining,  Mayer's  paracarmine  and  Mayer's  hemalum  are 
highly  recommended  by  nearly  all  specialists  on  these  forms.  The  animals 
should  be  much  overstained  and  then  very  rapidly  and  completely  destained 
in  strongly  acidulated  (2  to  4  per  cent  HC1)  70  per  cent  alcohol  (Cort, 
Transactions  of  the  American  Microscopical  Society,  XXXIV,  No.  4  [October, 
1916]). 

To  prevent  the  curling  up  of  flat  worms  which  are  to  be  infiltrated  for 
sectioning,  Peaslee  binds  them  by  wrappings  of  thread  to  a  bit  of  bristol 
board  which  is  not  removed  until  the  animal  is  to  be  imbedded. 

9.  Spirogyra,  Protococcus,  Volvox,  Desmids,  etc.,  may  be  mounted  in  a 
cell  in  the  following  copper  solution: 

Acetate  of  copper 1  gram 

Camphor  water 240     c.c. 

Glycerin 240     c.c. 

Glacial  acetic  acid ' 0.3  c.c. 

Corrosive  sublimate,  saturated  aqueous  solution . .  0 . 1  c.c. 


Objects  of  General  Interest  101 

Mix  thoroughly,  filter,  and  keep  in  a  glass-stoppered  bottle.  The  green 
color  of  the  plant  may  frequently  be  preserved  for  some  time  in  this  medium. 
The  specimen  is  washed  in  water,  transferred  to  the  cell,  then  the  solution 
is  added.  The  cell  is  covered  and  sealed  in  the  usual  way. 

10.  A  Dipping-Tube  is  a  simple  glass  tube.    To  operate  it,  hold  the 
tip  of  the  forefinger  over  the  upper  end  and  dip  the  lower  end  into  the  water 
until  it  comes  just  above  the  object  desired;  lift  the  finger  and  let  the  air 
out  of  the  tube,  and  the  water  will  rush  in  at  the  lower  end  carrying  the  object 
with  it.    Replace  the  ringer  over  the  top  of  the  tube  and  remove  it;   the 
water  will  remain  in  it  as  long  as  the  finger  is  held  firmly  over  the  upper  end. 
When  the  finger  is  removed  the  water  and  the  object  pass  out.    The  object 
may  sometimes  be  more  readily  discharged  if  the  tube  is  rotated.    A  pipette 
made  of  a  large-bore  glass  tube  and  an  atomizer  bulb  is  also  very  serviceable. 

1 1.  To  Keep  Water  from  Evaporating  from  a  Cell  Too  Freely,  use  a  round 
cell  and  cover  it  with  a  square  cover-glass.    Apply  a  brush  wet  with  water 
to  the  slide  beneath  one  of  the  projecting  corners  of  the  cover  from  time  to 
time.     Capillary  attraction  will  draw  in  the  water  and  will  keep  the  cell 
full.    If  a  continuous  supply  of  fresh  water  is  necessary,  one  end  of  a  loosely 
twisted  cotton  thread  may  be  laid  along  one  side  of  the  cover  and  the  other 
end  of  the  thread  immersed  in  a  small  vessel  of  water  which  stands  within 
half  or  three-quarters  of  an  inch  of  the  cell.    A  reservoir  made  from  the 
bottom  of  a  shell  vial  or  homeopathic  vial  answers  very  well;   it  may  be 
cemented  to  the  slide. 

Protozoa  and  other  small  forms  may  be  kept  alive  on  a  slide  for  a  number 
of  hours  by  simply  mounting  them  in  water  under  a  cover  in  a  cell  of  blotting 
paper  which  has  been  saturated  with  water.  For  aquaria  for  studying 
microscopic  organisms,  Walton  (Ohio  State  University  Bulletin,  XIX,  No.  5 
[1915])  used  ring-like  pieces  of  lens  paper  cut  somewhat  smaller  than  the 
cover-glass.  Such  aquaria  keep  for  several  hours.  They  may  be  made 
more  permanent  by  letting  them  stand  15  to  30  minutes  in  order  to  allow 
the  outside  water  to  evaporate,  and  then  running  paraffin  oil  around  the 
margin  of  the  cover-glass. 

12.  Deep  Cells  are  made  frequently  by  cutting  out  rings  of  paper,  lead, 
or  block-tin  with  gun  punches  and  cementing  them  to  the  slide.    Glass  and 
hard-rubber  rings  of  various  sizes  may  be  purchased  from  dealers.    To 
support  cover-glasses  Barker  uses  circular  cloth  patches  with  a  hole  in  the 
center.    These  may  be  bought  of  a" stationer. 

13.  Filtered  Shellac  is  recommended  by  McClung  as  excellent  both  for 
making  and  for  sealing  cells.     It  may  be  colored  with  Bismarck  brown  and 
similar  dyes.    Barker  uses  any  good  quality  of  enamel  paint. 

14.  For  a  Method  of  Preserving  Fine  Dissections  for  microscopic  study, 
as  opaque  objects,  see  memorandum  15,  p.  134. 


102  Animal  Micrology 

15.  The  Clearing  of  Total  Specimens  as  developed  by  Spalteholz  (  Ueber 
das  Durchsichtigmachen  von  menschlichen  und  tierschen  Praparaten,  2d  ed., 
1914,  S.  Hirzel,  Leipzig),  whereby  relatively  large  anatomical  and  embryo- 
logical  preparations  can  be  made  transparent,  is  an  extremely  useful  method, 
particularly  with  injected  objects.  The  following  account  of  the  method 
together  with  modifications  introduced  by  herself  is  taken  from  Miss  Sabin's 
article  in  Contributions  to  Embryology,  III,  No.  7,  Carnegie  Institution  of 
Washington,  1915:  "In  general  the  essentials  of  the  method  are,  first, 
fixation  in  formalin;  second,  a  thorough  bleaching  of  the  tissues  with  hydro- 
gen peroxide  to  remove  the  hemaglobin  and  other  pigments;  third,  dehydra- 
tion; and,  fourth,  clearing  the  specimens  in  an  oil  which  has  the  same  index 
of  refraction  as  the  tissues.  As  applied  to  embryonic  tissues,  the  method, 
developed  by  Professor  Spalteholz,  to  whom  I  am  very  much  indebted,  is 
as  follows:  The  specimens  which  have  been  injected  with  India-ink  are 
fixed  for  24  to  48  hours  in  5  and  10  per  cent  formalin.  Commercial  formalin 
is  slightly  acid,  which  is  an  advantage  for  the  India-ink  injections,  since  the 
ink  .diffuses  in  an  alkaline  solution.  Specimens  which  have  been  injected 
with  silver  nitrate  are  ruined  by  fixation  in  formalin,  because  the  silver  salt 
is  changed  to  a  white  precipitate  which  obscures  the  vessels.  If  injections 
of  bone  are  desired,  the  formalin  may  be  made  slightly  alkaline  and  the 
diffusion  of  the  ink  prevented  as  much  as  possible  by  tying  off  all  vessels 
before  fixation.  For  large  fetuses,  which  are  to  be  cleared  in  toto,  Dr.  P.  G. 
Shipley  has  found  that  the  subsequent  bleaching  is  made  easier  by  washing 
the  specimen  in  running  water  before  fixation,  thus  removing  much  of  the 
hemaglobin.  After  fixation,  the  specimens  are  washed  in  running  tap  water 
from  12  to  24  hours,  followed  by  distilled  water  to  remove  the  formalin. 
The  bleaching  is  done  in  hydrogen  peroxide.  Spalteholz  adds  a  few  drops 
of  ammonia  to  precipitate  the  barium  salts.  This  is  not  necessary  with 
barium-free  oxide.  For  adult  tissues,  Spalteholz  uses  undiluted  peroxide; 
for  the  embryonic  tissues  about  2  to  3  per  cent  is  the  best  strength.  The 
small  embryos  with  ink  injections  take  about  20  minutes  to  bleach;  for  the 
silver  specimens,  2  to  3  minutes  suffice,  and  they  must  be  watched  constantly 
and  the  bleaching  stopped  before  the  silver  is  affected.  Following  the 
bleaching,  the  specimens  must  be  washed  thoroughly  in  running  water  and 
in  distilled  water.  The  dehydration  may  be  begun  with  50  per  cent  alcohol 
and  the  percentage  increased  successively  by  five  points  or  less.  After  two 
changes  of  a  good  grade  of  absolute  alcohol,  the  specimens  are  passed  through 
changes  of  benzene  into  the  synthetic  oil  of  wintergreen.  The  small  amount 
of  benzene  which  is  carried  over  evaporates  quickly,  and  the  few  bubbles 
which  develop  in  the  bleaching  process  can  be  removed  with  needles.  The 
oil  of  wintergreen  should  be  entirely  colorless,  but  both  the  specimens  and 
the  oil  will  gradually  become  brown  with  age.  This  is  especially  true  of  the 


Objects  of  General  Interest  103 

silver-nitrate  specimens,  but  they  will  keep  for  six  months  or  a  year  in  oil. 
They  can  be  returned  to  alcohol  for  storage  and  recleared  when  desired,  or 
they  may  be  made  permanent  in  balsam.  The  advantage  of  keeping  the 
total  specimens  in  oil  rather  than  in  balsam  is  that  they  can  be  dissected. 
On  the  other  hand,  they  are  made  more  permanent  in  balsam.  The  oil  of 
wintergreen  makes  the  tissues  tough,  so  that  it  is  possible  to  obtain  minute 
dissections  of  the  injected  specimens. 

"The  Spalteholz  method  as  applied  to  embryos  can  be  very  much 
simplified  by  changing  the  fixative.  For  mammalian  embryos  the  best 
fixative  is  Carney's  mixture.  This  is  absolute  alcohol  60  parts,  chloroform 
30  parts,  and  glacial  acetic  acid  10  parts.  In  this  mixture  the  acid  is  suffi- 
ciently strong  to  bleach  the  hemaglobin  so  that  the  peroxide  is  unnecessary. 
The  penetrating  power  of  the  fixative  is  very  great,  which  is  of  importance, 
since  no  injected  specimen  can  be  cut  into  until  it  is  thoroughly  fixed.  The 
relations  of  the  tissues  are  well  maintained  and  the  swelling  due  to  the  acetic 
acid  tends  to  counteract  the  shrinkage  that  always  takes  place  in  the  oil  of 
wintergreen.  The  fixative  does  not  affect  any  of  the  injection  fluids.  The 
process  after  fixation  in  the  Carney's  mixture  is  simple;  specimens  remain 
in  the  fixative  from  2  to  12  hours  and  are  then  placed  directly  into  70  per 
cent  alcohol,  dehydrated  in  graded  alcohols,  and  cleared  as  before.  The 
specimens  can  then  be  studied  in  toto,  or  dissected  or  imbedded  in  paraffin 
and  sectioned.  They  should  be  imbedded  through  a  mixture  of  the  oil  of 
wintergreen  and  paraffin.  They  do  not  become  brittle  in  the  oil,  so  that  they 
may  be  sectioned  after  staying  in  the  oil  for  many  weeks.  The  shrinkage 
in  the  oil,  however,  seems  to  increase  on  long  standing.  The  advantages 
of  the  fixation  hi  Carney's  mixture  are  that  the  specimens  are  even  clearer 
than  after  bleaching  with  peroxide,  there  are  no  bubbles  formed  to  damage 
the  tissues,  the  time  of  the  procedure  is  shortened,  and  the  fixation  is  much 
better  should  it  be  desired  to  section  the  specimens  after  studying  the  vessels 
in  whole  embryos.  Specimens  which  are  strongly  pigmented,  however, 
must  be  bleached  with  hydrogen  peroxide  before  they  can  be  cleared." 

For  other  methods  of  clearing  in  toto  preparations  see  memoranda  16 
and  17. 

16.  The  Potash  Clearing  Method  (or  Modifications  of  It)  for  "in  Toto" 
Preparations  is,  according  to  Thurlow  C.  Nelson  of  our  own  laboratories, 
one  of  the  best  as  well  as  one  of  the  simplest  methods  for  the  demonstration 
of  skeletal  and  cartilaginous  structures.  Unless  nervous  or  other  structures 
are  to  be  stained,  the  animal  should  be  put  into  a  1  per  cent  potash  solu- 
tion immediately  after  killing.  Twenty-four  hours  in  this  medium  should 
be  sufficient  to  clear  the  overlying  tissues  to  such  an  extent  that  the  skeletal 
elements  are  clearly  visible.  After  removal  to  glycerin  the  specimen  will 
keep  indefinitely. 


104  Animal  Micrology 

For  Staining  Nerve  Tissue  "in  Toto,"  Nelson  combines  the  Charles 
Sihler  hematoxylin  stain  with  the  potash  method.  Three  solutions  are 
required: 

1 
Potassium  hydroxide,  1  per  cent  aqueous  solution 

2 

Glacial  acetic  acid 1  part 

Glycerin 1  part 

Chloral  hydrate,  1  per  cent  solution 6  parts 

3 

Glycerin 1  part 

Ehrlich's  acid  hematoxylin 1  part 

Chloral  hydrate,  1  per  cent  solution 6  parts 

A  minnow,  for  example,  is  killed  in  95  per  cent  alcohol  and  left  for  48 
hours.  After  the  viscera  are  removed,  it  is  next  transferred  to  the  potash 
solution  for  from  1  to  3  days.  When  transparent,  the  specimen  is  put  into 
solution  No.  2  for  72  hours,  then  into  solution  No.  3  for  a  week.  It  is  then 
destained  in  solution  No.  2  for  18  hours  and  cleared  in  glycerin.  The 
nervous  tissue  should  show  up  dark  purple  in  the  semi-transparent  muscular 
tissue  of  the  body. 

17.  A  Benzaldehyde  Clearing  and  Fixing  Method  for  "in  Toto"  Prepara- 
tions has  also  been  devised  by  Nelson  (memorandum  16)  as  follows: 

Small  objects,  such  as  eggs  and  small  fish,  may  be  put  into  the  aldehyde 
directly  as  it  acts  as  a  dehydrating  agent  to  a  slight  extent,  being  miscible 
with  30  parts  of  water.  Larger  objects  are  dehydrated  by  running  up 
through  the  alcohols  and  then  clearing  in  the  benzaldehyde.  If  a  specimen 
of  the  fresh-water  mussel,  for  example,  Anodon,  is  treated  with  50  per  cent 
alcohol  for  2  hours,  70  per  cent  for  5  hours,  80  per  cent  for  15  hours,  and 
95  per  cent  for  2  hours,  and  is  then  put  into  benzaldehyde,  the  mantle  begins 
to  clear  at  once,  the  whole  preparation  being  almost  transparent  in  12  hours. 
As  benzaldehyde  is  very  unstable,  being  oxidized  to  form  benzoic  acid,  it 
must  be  removed  before  the  tissue  is  exposed  to  the  air  for  any  long  period. 
Likewise  vessels  containing  it  must  be  kept  tightly  stoppered.  Objects 
may  be  kept  in  the  fluid  permanently  by  sealing  the  container  in  such  a  way 
as  to  exclude  all  air.  This  method  of  clearing  finds  its  chief  use  in  clearing 
up  the  entire  bodies  of  small  animals  so  as  to  show  injected  circulatory 
systems  and  such  structures.  Small  objects  may  be  mounted  in  balsam 
from  benzaldehyde  if  preferred. 

To  infiltrate  with  paraffin  or  to  stain  in  toto,  the  object  should  first  be 
passed  through  a  mixture  of  6  parts  of  xylol  to  1  part  of  absolute  alcohol. 
A  satisfactory  stain  may  be  prepared  by  dissolving  methylen-blue  crystals 
in  2  c.c.  of  absolute  alcohol  and  adding  6  c.c.  of  benzene. 


CHAPTER  XIV 

BLOOD 

I.    EXAMINATION  OF  FRESH  BLOOD 

a)  General. — 1.  Thoroughly  clean  a  slide  and  cover,  bathe  one  thumb 
in  ether-alcohol  (reagent  4,  p.  8),  sterilize  a  sharp  needle  by  heating  it 
in  a  flame,  and  then  prick  the  back  of  the  thumb  with  the  needle. 

2.  Place  a  small  drop  of  the  resulting  blood  on  a  slide  and  quickly  put 
on  a  cover-glass.  To  prevent  evaporation,  the  edges  of  the  cover  may  be 
surrounded  by  olive  oil  or  vaseline. 

Living  corpuscles  may  also  be  studied  in  a  drop  of  normal  saline  or  in 
Ringer's  solution  (p.  239). 

b)  Effects  of  reagents. — When  it  is  desired  to  study  the  effects  of  reagents 
on  fresh  blood  (e.g.,  distilled  water,  1  per  cent  tannic  acid,  etc.),  a  drop  of 
fresh  blood  is  placed  on  a  slide,  the  cover  is  put  on,  and  then  the  blood  is 
"irrigated "  with  the  reagent.     That  is,  a  drop  of  the  reagent  is  placed  at  the 
edge  of  the  cover  to  be  drawn  under  .by  capillary  action.     The  process  may 
be  hastened  by  applying  the  edge  of  a  bit  of  blotting  paper  to  the  opposite 
edge  of  the  cover. 

c)  To  demonstrate  blood-platelets. — Place  a  small  drop  of  a  1  per  cent 
solution  of  methyl  violet  (reagent  61,  p.  231)  in  normal  salt  solution,  on  the 
back  of  a  thumb  which  has  been  cleaned  by  washing  it  in  ether-alcohol. 
With  a  sterilized  needle  prick  the  thumb  through  the  stain  and  mount  a 
drop  of  the  blood  which  exudes.     Examine  it  under  a  high  power.     Both 
platelets  and  white  corpuscles  are  stained. 

d)  Stained  preparation  of  fibrin. — Mount  a  drop  of  blood  on  a  slide  as  in 
a.     Place  it  in  a  moist  chamber  for  from  20  to  30  minutes  to  coagulate. 
Loosen  the  cover  with  a  few  drops  of  water  and  then  thoroughly  irrigate  the 
preparation  with  water.     Drain  off  the  water,  blot  the  preparation  with 
blotting  paper,  and  add  immediately  a  drop  of  a  1  per  cent  aqueous  solution 
of  eosin  (reagent  43,  p.  224) .     Remove  this  after  3  minutes,  rinse  the  prepara- 
tion in  water,  then  treat  it  3  minutes  with  a  1  per  cent  aqueous  solution  of 
methyl  violet  (reagent  61,  p.  231).     Rinse  the  preparation  in  water,  let  it 
dry,  and  finally  mount  in  balsam. 

e)  Crystals  of  the  blood. — 

1.  Hemoglobin  Crystals. — Allow  a  drop  of  blood  to  dry  on  the  slide 
without  covering  it.  Long  rhombic  prisms  of  a  red  color  crystallize  out. 
The  blood  of  a  rat  is  best  for  demonstration.  A  more  certain  method  is  as 

105 


106  Animal  Micrology 

follows:  To  5  c.c.  of  blood  in  a  test-tube  add  a  few  drops  of  ether  and  shake 
the  mixture  vigorously  until  the  blood  becomes  laky.  Place  a  drop  or  two 
of  the  laked  blood  on  a  slide  and  allow  it  to  dry  in  the  cold. 

2.  Hematoidin  Crystals;    reddish-yellow  crystals  (rhombic  plates). — 
They  can  be  obtained  from  old  blood  extravasations  (e.g.,  cerebral  hemor- 
rhage, corpora  lutea,  etc.)  by  teasing.    Mount  in  Canada  balsam. 

3.  Hemin  or  Teichmann's  Crystals. — To  a  small  drop  of  blood  on  a  siido 
or  a  bit  of  cloth  which  has  been  previously  saturated  with  blood,  add  a 
few  crystals  of  common  salt.    Heat  over  a  flame  until  the  mixture  has 
become  dry,  leaving  a  reddish-brown  residue.    Apply  a  cover-glass  and 
flood  the  preparation  with  as  much  acetic  acid  as  will  remain  in  place  under 
the  cover.    Heat  the  preparation  until  the  acetic  acid  boils.    After  the 
acid  has  evaporated,  the  preparation  may  be  made  permanent  by  adding 
Canada  balsam.    The  crystals  are  very  small,  narrow  rhombic  plates  of 
dark-brown  color.    They  vary  in  size  and  may  lie  singly,  across  one  another, 
or  in  stellate  groups. 

The  presence  of  these  crystals  is  positive  evidence  of  the  presence  of 
blood,  hence  their  demonstration  is  of  great  importance  in  stains  or  fluid 
suspected  of  containing  blood. 

II.    COVER-GLASS  PREPARATIONS 

a)  Dry  preparations  (Ehrlich's  method). — 1.  In  this  method  the  prepara- 
tion is  "fixed"  by  means  of  heat.  Under  one  end  of  a  copper  bar  or  copper 
triangle  (Fig.  26)  place  a  flame.  After  15  or  20  minutes  a  given  point  on 
the  bar  will  have  a  practically  constant  temperature.  Thoroughly  clean  the 
bar,  run  a  stream  of  water  along  the  top  of  it  toward  the  flame,  and  locate 
the  point  farthest  from  the  flame  at  which  the  water  boils.  The  blood 
smears  when  prepared  are  to  be  placed  film  side  up  in  a  row  across  the  bar 
about  three-fourths  of  an  inch  nearer  the  flame  than  the  point  at  which  the 
water  just  boiled.  This  will  subject  them  to  a  temperature  of  about  120°  C. 

2.  Thoroughly  clean  and  dry  two  cover-glasses,  touch  one  to  a  small  drop 
of  perfectly  fresh  blood  as  it  comes  from  the  finger  or  lobe  of  the  ear  and 
instantly  drop  it  on  to  the  second  cover.  The  blood  should  spread  in  a 
thin  film  between  the  covers;  if  it  does  not,  it  has  begun  to  coagulate  and 
the  preparation  will  be  inferior.  Rapidly  separate  the  covers  by  sliding  them 
apart,  wave  them  in  the  air  a  minute  to  dry  the  films,  then  place  them  down 
with  the  smear  side  uppermost.  Do  not  press  the  covers  together  to  spread  the 
blood  because  this  ruins  the  corpuscles.  If  the  red  corpuscles  are  to  retain 
their  shape  the  film  of  blood  must  be  extremely  and  uniformly  thin.  Practice 
until  you  have  prepared  such  a  film. 

Some  workers  prefer  to  make  smears  on  slides  instead  of  on  cover-slips. 
A  small  drop  of  blood  placed  near  one  end  of  a  perfectly  clean  slide  is  spread 


Blood  107 

by  drawing  an  edge  of  the  end  of  another  slide  through  the  drop  and  along 
the  surface  of  the  first  slide  to  its  opposite  end.     Crushing  is  thus  avoided. 

NOTE. — In  the  clinical  examination  of  blood  great  care  must  be  exercised 
to  have  it  absolutely  fresh;  furthermore,  the  cover-glasses  should  be  handled 
with  forceps  instead  of  by  means  of  the  fingers.  It  is  recommended  that  one  pair 
of  the  forceps  be  Coronet  or  spring  forceps  of  some  kind  (Fig.  39).  The  lobe  of 
the  ear  is  perhaps  the  best  region  from  which  to  obtain  the  blood.  The  needle 
with  which  the  puncture  is  made  should  always  be  sterilized.  Wipe  away  the 
first  drop  of  blood  that  appears.  The  drop  finally  chosen  should  be  one  that 
has  appeared  immediately  after  the  spot  has  been  wiped  and  it  should  be  but 
little  larger  than  a  pin-head.  The  whole  operation  cannot  be  performed  too 
rapidly.  To  shorten  the  time  it  is  well  to  have  an  assistant  to  prick  and  manipu- 
late the  ear  while  the  operator  attends  to  the  preparation  of  the  film. 

3.  When  several  satisfactory  films  have  been  prepared,  place  them  on 
the  heated  bar,  as  indicated  in  step  1.    Cover  them  to  keep  out  dust  and  leave 
them  for  from  30  to  60  minutes. 

4.  Remove  the  covers  and  stain  the  preparations  15  or  20  minutes  with 
Ehrlich's  triple  stain  (reagent  42,  p.  223)  by  flooding  the  film  with  the  stain 
by  means  of  a  pipette.     Rinse  off  the  surplus  stain  with  water,  blot  the  film 
with  blotting  paper,  and  dry  it  by  holding  it  with  the  edge  downward  high 
above  the  flame.    When  dry,  mount  in  balsam  on  a  slide. 

NOTE. — Instead  of  heating  the  preparation,  much  the  same  results  may  be 
obtained  by  subjecting  films  (prepared  as  in  step  2)  to  ether  and  alcohol  (p.  8) 
for  from  1  to  12  hours,  drying  them  again  in  the  air,  and  then  staining  as  above. 

6)  Rapid  method. — 1.  Prepare  a  film  as  above  (a,  2),  but  before  it  has 
dried  treat  it  for  30  minutes  with  a  saturated  aqueous  solution  of  corrosive 
sublimate  (reagent  14,  p.  212). 

2.  Wash  the  preparation  thoroughly  in  water  or  in  50  per  cent  alcohol. 

3.  Stain  for  10  minutes  in  Delafield's  or  Ehrlich's  hematoxylin  (reagent 
52  or  53,  p.  226),  rinse  in  70  per  cent  alcohol,  and  stain  for  20  seconds  hi 
eosin  (0 . 5  per  cent  solution  in  70  or  95  per  cent  alcohol). 

4.  Rinse  in  95  per  cent  and  in  absolute  alcohol  each  for  2  minutes,  pass 
through  xylol,  and  mount  in  balsam. 

After  rinsing  following  staining,  some  workers  simply  blot  the  prepara- 
tion with  blotting  paper,  dry  it  in  the  air,  and  mount  it  in  balsam. 

m.    ENUMERATION  OF  BLOOD  CORPUSCLES 

The  instrument  used  is  the  hemocytometer  (Fig.  38).  It  consists  of  a 
special  slide  for  counting  and  two  graduated  pipettes  for  diluting  and  measur- 
ing the  blood. 

Obtain  a  drop  of  blood  from  the  lobe  of  the  ear  or  from  the  finger.  Fill 
the  smaller  pipette  (with  the  101  mark  just  above  the  chamber  containing 


108 


Animal  Micrology 


bead)  of  the  hemocytometer  to  the  mark  1  by  careful  suction.  The  tip  of 
the  tongue  placed  firmly  over  the  hole  in  the  mouthpiece  will  keep  the 
blood  from  dropping  back.  If  the  blood  is  drawn  beyond  the  1  mark,  blow 
it  out  immediately,  clean  the  tube,  and  repeat  the  operation. 

Wipe  the  blood  from  the  outside  of  the  pipette  quickly  and  draw  in 
sufficient  Toisson's  solution  to  make  the  level  of  the  combined  liquids  stand 
precisely  at  the  mark  101.  Close  the  ends  of  the  pipette  with  thumb  and 


FIG.  38. — Hemocytometer 

a,  view  of  slide  from  above;  6,  view  of  slide  from  one  side;  c,  counting-disk  which 
lies  at  the  center  of  B;  E,  bead  for  mixing;  M,  mouthpiece. 

middle  finger  and  mix  the  blood  thoroughly  with  the  solution  by  shaking 
the  tube  for  2  minutes.     The  blood  is  thus  diluted  100  times. 

Toisson's  solution: 

Sodium  sulphate 8.0      grams 

Sodium  chloride 1.0      gram 

Neutral  glycerin 30          c.c. 

Methyl  violet,  5b 0 . 025  gram 

Distilled  water 160          c.c. 

Blow  out  a  drop  of  the  liquid  to  remove  the  unmixed  solution  remaining 
in  the  capillary  tube.  Have  the  counting-disk  and  cover-glass  perfectly 
clean.  Allow  a  drop  of  the  diluted  blood  to  flow  on  to  the  disk  and  place 
the  cover-glass  over  the  drop.  The  cell  of  the  disk  must  be  entirely  filled 
by  the  drop  of  blood.  Let  the  corpuscles  settle  a  minute  or  two  before 
beginning  to  count.  Examine  the  field  under  a  low-power  objective  to  see 
that  the  corpuscles  are  evenly  distributed.  If  they  are  not,  the  blood  was 
not  thoroughly  mixed  and  the  whole  operation  should  be  repeated  after 
thoroughly  cleansing  the  pipette. 

Examine  the  preparation  under  a  high  power  of  the  microscope,  and 
count  the  number  of  red  corpuscles  in  20  to  40  small  squares;  of  those 
corpuscles  whJch  happen  to  lie  on  the  boundary  line,  count  the  ones  that  lie 


Blood  109 

only  in  the  upper  and  on  the  left  sides  of  each  square.  Take  the  average 
number  in  a  square  and  calculate  the  number  of  corpuscles  in  a  cubic  milli- 
meter of  blood. 

The  depth  of  the  entire  cell  is  0. 1  mm.,  the  area  of  each  small  square 
is  T^TTF  sq.  mm.,  consequently  the  volume  of  blood  in  each  square  column  is 
ToW  cu.  mm.,  or  1  cu.  mm.  of  diluted  blood  would  contain  4,000  times  the 
average  number  in  a  square.  One  cubic  millimeter  of  undiluted  blood  con- 
tains 100  times  as  many,  or  400,000  times  the  number  in  one  square.  What 
result  do  you  obtain  ?  For  accuracy,  three  separate  counts  should  be  made 
and  the  average  taken. 

After  finishing  the  count,  clean  the  pipette  by  successively  drawing 
into  and  expelling  from  it  water,  alcohol,  and  finally  ether.  Do  not  blow 
through  it,  but  cause  the  ether  to  evaporate  by  sucking  air  through  the  tube. 
For  counting  the  white  corpuscles  use  the  large  pipette  and  dilute  the  blood 
10  times  with  one-third  of  1  per  cent  glacial  acetic  acid.  The  acid  destroys 
the  red  corpuscles,  and  thus  the  white  corpuscles  are  more  readily  seen. 
Proceed  in  the  same  manner  as  for  red  corpuscles. 

Use  water  instead  of  alcohol  for  cleaning  the  counting-slide,  as  alcohol 
dissolves  the  cement  of  the  slide. 

IV.     OBSERVATION  OF  THE  BLOOD  CURRENT 

a)  Circulation  in  the  web  of  a  frog's  foot. — Wind  a  long  strip  of  cheese- 
cloth around  a  frog  stretched  out  upon  a  narrow  piece  of  thin  board,  leaving 
one  hind  foot  exposed.    Soak  the  cloth  in  water  in  order  to  keep  the  animal's 
skin  moist.    Pin  the  extended  foot  in  such  a  way  that  the  web  between  the 
toes  is  stretched  over  a  notch  or  hole  in  the  end  of  the  board.    Examine  under 
the  microscope.    If  the  preparation  is  favorable,  leucocytes  may  perhaps 
be  seen  penetrating  the  walls  of  the  vessel  (diapedesis)  and  passing  into  the 
surrounding  tissues. 

b)  Circulation   in   the   mesentery.    Inflammation. — Immobilize    a   frog 
(the  male  is  better)  by  injecting  a  few  drops  of  a  1  per  cent  solution  of  curare 
into  one  of  the  dorsal  lymph  sacs.    Curare  paralyzes  the  nerve-endings. 
After  waiting  20  minutes  for  the  curare  to  be  absorbed  into  the  circulation, 
cut  open  the  abdominal  wall  for  a  short  distance  along  the  left  side  and  draw 
out  several  loops  of  the  intestine.    Pin  out  a  favorable  area  of  mesentery 
over  a  cork  ring,  and,  after  covering  it  with  a  cover-glass,  examine  under  the 
microscope.    Keep  the  parts  moistened  with  normal  salt  solution.    Such  a 
preparation  is  especially  favorable  for  studying  the  migrations  of  leucocytes 
through  the  walls  of  the  vessels.    Do  not  have  the  mesentery  stretched  too 
tightly  or  the  circulation  will  cease.    After  a  time  the  phenomena  of  inflam- 
mation may  readily  be  observed.    It  is  hastened  if  some  irritant  (e.g.,  a 
drop  of  creosote)  is  applied  to  the  mesentery. 


110  Animal  Micrology 

MEMORANDA 

1.  For  Demonstration  of  the  Different  Granules  of  Leucocytes,  etc.,  see 
p.  240,  under  the  general  topic  of  blood. 

2.  To  Study  Blood  in  Sections,  ligate  a  small  vessel  in  two  places  to 
keep  in  the  corpuscles,  then  remove  the  piece  so  prepared  and  fix  it  in  Bouin's 
fluid  or  osmic  acid.    Imbed  in  paraffin  and  cut  thin  sections.    Stain  material 
fixed  in  Bouin  or  corrosive-sublimate  reagents  by  the  hematoxylin-eosin 
method  (p.  50)  or  with  the  Ehrlich-Biondi  stain  (41,  p.  222).    The  blood 
fixed  in  osmic  acid  may  be  stained  by  the  saffranin-gentian  violet  method 
(73,  p.  234). 

3.  Amoeboid  Movements  in  Leucocytes  may  readily  be  observed  in 
blood  (preferably  amphibian)  which  has  been  mounted  on  a  slide  in  very 
slightly  warmed  normal  saline.    Place  a  hair  under  the  cover-glass  and  seal 
the  edges  of  the  latter  with  vaseline  or  melted  paraffin.    For  continuous 
study  of  the  white  corpuscles  of  warm-blooded  animals  a  warm  stage  of 
some  kind  is  necessary  to  keep  the  temperature  of  the  blood  near  the 
temperature  of  the  body. 

4.  Ingestion  by  Leucocytes. — Rub  up  sufficient  India  ink  in  a  few  drops 
of  normal  saline  to  make  a  grayish  fluid.    With  fine  scissors  make  an  inci- 
sion into  one  of  the  dorsal  lymph  sacs  of  a  chloroformed  frog  (parallel  to  and 
close  beside  the  urostyle) .    Introduce  a  capillary  pipette  into  the  wound  and 
obtain  a  small  drop  of  lymph.    Mix  it  on  a  slide  with  a  drop  or  two  of  the 
prepared  ink.    After  placing  a  hair  across  the  field,  put  on  a  cover-glass  and 
seal  the  edges  with  vaseline  or  melted  paraffin.    Under  a  high  power  of  the 
microscope  the  cells  may  be  seen  engulfing  the  colored  particles. 

Gage  (The  Microscope)  recommends  a  mixture  of  lamp-black,  2  grams; 
sodium  chloride,  1  gram;  gum  Arabic,  1  gram;  distilled  water,  100  c.c. 
Mix  thoroughly  in  a  mortar  and  filter  through  one  layer  of  gauze  and  one 
of  lens  paper.  When  injected  into  an  animal  the  leucocytes  will  ingest  the 
particles  of  carbon. 

5.  Wright's  Stain  for  Blood. — This  is  a  modification  of  Irishman's 
Romanowsky  stain.    To  prepare  the  stain  make  a  0 . 5  per  cent  solution  of 
sodium  bicarbonate  in  distilled  water  and  add  to  it  1  per  cent  of  methylen 
blue  (B.X.,  or  "medicinally  pure"):    Subject  the  mixture  to  live  steam 
in  an  ordinary  steam  sterilizer  (e.g.,  Arnold;  not  a  pressure  sterilizer  or  a 
water-bath)  for  one  hour.    The  container  should  be  of  such  size  that  the 
liquid  forms  a  layer  not  more  than  6  cm.  deep.    When  the  mixture  is  cool, 
filter  to  remove  any  precipitate.    To  each  100  c.c.  of  the  filtered  mixture 
add,  with  constant  stirring,  500  c.c.  of  a  0.1  per  cent  aqueous  solution  of 
"yellow,  water-soluble"  eosin.     Collect  the  resulting  precipitate  on  a  filter, 
dry  it  thoroughly,  and,  rubbing  up  in  a  porcelain  dish  or  mortar  if  necessary, 
make  a  5  per  cent  solution  in  pure  methylic  alcohol.    To  prevent  the  alcohol 


Blood  111 

from  evaporating,  keep  the  bottle  containing  the  solution  tightly  stoppered. 
Should  precipitation  occur,  filter  the  stain  and  add  a  small  quantity  of  methyl 
alcohol. 

Mallory  and  Wright  in  their  Pathological  Technique,  p.  383,  give  the 
following  si.innTnfl.ry  of  the  method  for  staining  blood  films: 

1.  Make  films  of  the  blood,  spread  thinly,  and  allow  them  to  dry  in 
the  air. 

2.  Cover  the  preparation  with  a  measured  quantity  of  the  staining  fluid 
for  one  minute. 

3.  Add  to  the  staining  fluid  on  the  preparation  the  same  quantity  of 
distilled  water  as  there  was  of  the  stain.    Allow  this  mixture  to  remain  on 
the  preparation  for  two  or  three  minutes,  according  to  the  intensity  of  the 
staining  desired.    Eosinophilic  granules  are  best  brought  out  by  briefer 
staining. 

4.  Wash  in  water,  preferably  in  distilled  water,  until  the  film  has  a 
pinkish  tint  in  its  thinner  or  better-spread  portions  and  the  red  corpuscles 
acquire  a  yellow  or  pink  color. 

5.  Dry  between  filter-paper  and  mount  in  balsam.    The  preparations 
retain  their  colors  as  long  as  any  preparations  stained  with  anilin  dyes. 
Fresh  films  stain  better  than  those  which  are  several  hours  old. 

Erythrocytes  when  stained  by  Wright's  stain  should  appear  orange  or 
pink  in  color  (with  deep-blue  nuclei,  when  nucleated) ;  lymphocytes  should 
show  purplish-blue  nuclei  and  cytoplasm  of  robin's-egg  blue  with  occasional 
dark-blue  or  purplish  granules;  polynuckar  neutrophilic  leucocytes  should 
have  blue  or  dark  lilac-colored  nuclei  with  cytoplasmic  granules  of  reddish- 
lilac  color;  eosinophilic  leucocytes  should  show  blue  or  dark  lilac-colored 
nuclei,  and  blue  cytoplasm  with  granules  the  color  of  eosin;  large  mononuckar 
leucocytes  should  show  blue  or  dark  lilac-colored  nuclei  with  cytoplasm 
pale  blue  in  one  form  and  blue  with  dark  lilac  or  deep  purple-colored  granules 
in  the  other;  mast-cells  should  exhibit  irregular-shaped  nuclei  stained 
purplish  or  dark-blue  in  bluish  cytoplasm  in  which  numerous  coarse  spherical 
granules  of  variable  size,  dark  purple  to  black  in  color,  are  imbedded;  myelo- 
cytes  have  dark-blue  or  dark  lilac-colored  nuclei  and  blue  cytoplasm  con- 
taining numerous  dark-lilac  or  reddish-lilac-colored  granules;  blood-platekts 
are  stained  blue. 

6.  For  Malarial  Parasites  Wright's  stain  (memorandum  5)  is  excellent. 
It  yields  the  so-called  Romanowsky  stain;    the  color  of  the  chromatin 
varies  from  lilac  to  very  dark  red,  while  the  body  of  the  parasite  stains 
blue.    A  full  account  of  the  method  will  be  found  in  Mallory  and  Wright's 
Pathological  Technique,  p.  447. 

7.  Ehrlich's  Triple  Stain  for  blood  is  given  on  p.  223. 


CHAPTER  XV 

BACTERIA 

No  attempt  is  made  here  to  give  even  an  elementary  account  of  bac- 
teriological technique.  Only  such  phases  of  the  work  as  are  concerned  with 
the  immediate  microscopical  examination  of  bacteria  are  touched  upon,  and 
these  chiefly  to  afford  some  practice  in  this  kind  of  manipulation.  For 
special  technique,  identification,  or  descriptions  of  apparatus  and  accessories, 
the  student  is  referred  to  standard  textbooks. 

BACTERIAL  EXAMINATION 

Bacteria  when  prepared  for  microscopical  examination  are  in  the  form  of 

A.  Cover-glass  preparations, 

B.  Bacteria  in  tissues  (section  method),  or 

C.  Hanging-drop  preparations. 

A.     Cover-Glass  Preparations 

I.   Killing  and  fixing. — 

1.  From  Fluid  Media  (e.g.,  bouillon,  milk,  water,  saliva,  blood,  pus, 
etc.) . — Sterilize  a  platinum  wire  loop  by  heating  it  red  hot  in  a  flame.  When 
cool,  touch  the  loop  to  the  culture  and  spread  the  adherent  bacteria  in  a 
thin  film  over  the  surface  of  a  cover-glass  which  has  been  sterilized  in  a 
flame.  After  the  film  has  dried  in  the  air,  kill  and  fix  the  bacteria  to  the  cover 
by  passing  it  three  times,  film  side  uppermost,  through  the  apex  of  a  flame. 


FIG.  39. — Cornet's  Cover-Glass  Forceps 

Each  time  should  not  exceed  half  a  second.  Prepare  several  films  from  a 
given  material.  Coronet  or  similar  forceps  (Fig.  39)  should  be  used  for 
handling  such  films,  because  the  cover-glass  can  be  left  in  them  through  the 
entire  operation  of  fixing  and  staining. 

If  a  platinum  loop  is  not  at  hand  a  second  cover-glass  may  be  used  to 
spread  the  smear.    The  first  cover-glass  is  held  in  a  pair  of  cover-glass 

112 


Bacteria  113 

forceps  and  the  second  cover-glass  is  dropped  on  to  it.  The  glasses  are 
then  rapidly  drawn  apart  with  a  sliding  motion  by  means  cf  forceps.  The 
glasses  should  not  be  pressed  tightly  together.  Proficiency  in  making  such 
preparations  is  gained  only  after  considerable  practice.  The  chief  secret 
in  making  a  good  preparation  is  to  get  the  films  extremely  thin  and  evenly 
distributed. 

2.  From  Solid  Media  (gelatin,  agar;  meat,  potato,  animal  tissues  and 
organs,  etc.). — The  procedure  is  the  same  as  for  1,  except  that  a  drop  of 
sterilized  water  or  bouillon  is  put  on  the  cover-glass  to  facilitate  the  spreading 
of  the  bacteria  in  a  film  over  the  cover. 

II.  Staining  and  mounting. — 

1.  Gentian  violet  (memorandum  3,  a,  p.  116),  5  minutes.    The  cover- 
glass  is  left  in  the  forceps,  film  side  up,  and  the  film  flooded  with  the  staining 
fluid. 

2.  Rinse  in  water. 

3.  Gram's  solution  (memorandum  3,  /,  p.  117)  until  the  color  becomes 
black  (2  to  3  minutes). 

4.  Ninety-five  per  cent  alcohol  until  the  violet  color  has  almost  com- 
pletely disappeared. 

5.  Rinse  in  water  and  examine  by  placing  the  cover-glass  film  side 
downward  on  a  slide.    Only  a  thin  film  of  water  should  remain  between 
the  slide  and  the  cover.    Remoye  surplus  water  by  means  of  blotting  paper. 
If  a  prolonged  examination  is  to  be  made,  water  lost  by  evaporation  must 
be  replaced  by  occasionally  placing  a  small  drop  of  water  at  the  edge  of  the 
cover.    In  ordinary  work  the  final  inspection  is  frequently  made  at  this  stage. 
If  a  permanent  preparation  is  desired,  however,  proceed  with  the  following 
steps: 

6.  If  the  bacteria  are  well  stained,  a  counterstain  of  Bismarck  brown 
(memorandum  3,  d,  solution  2,  p.  116)  may  be  added  (5  to  10  seconds).    This 
step  may  be  omitted. 

7.  Absolute  alcohol,  10  to  15  seconds. 

8.  Xyol. 

9.  Xylol-balsam. 

NOTE. — In  staining,  if  the  cover-glass  is  warmed  over  a  flame  some  15  or  20 
seconds  until  the  stain  steams,  the  action  of  the  stain  is  usually  more  intense 
and  more  rapid.  Boiling,  however,  must  be  avoided. 

B.    Bacteria  in  Tissues 

Tissues  may  be  fixed  and  hardened  (e.g.,  Gilson's  fluid,  Appendix  B, 
p.  213,  reagent  16;  or  Zenker's,  reagent  6;  or  formalin,  reagent  18)  in  the 
ordinary  way  and  sections  made  by  the  usual  methods.  Where  practicable, 
paraffin  sections  are  preferable  to  celloidin  sections,  because  the  celloidin 


114  Animal  Micrology 

tends  to  hold  the  stain  and  thus  obscure  the  bacteria.  Sections  should 
be  fixed  to  the  slide  (paraffin  by  albumen  fixative,  celloidin  by  ether 
vapor) . 

Bacteria  which  do  not  stain  by  the  Gram  method  (memorandum  3,  /, 
p.  117)  or  the  tubercle-bacillus  method  (memorandum  3,  e,  p.  117)  are  difficult 
to  demonstrate,  because  it  is  hard  to  stain  them  so  as  to  differentiate  them 
from  the  tissues  in  which  they  lie;  furthermore,  most  of  them  easily  lose 
whatever  stain  they  may  have  taken  up.  LofHer's  alkaline  methylen  blue 
(memorandum  3,  6,  p.  116)  is,  perhaps,  the  most  useful  stain  for  these  organ- 
isms. 

Methylen-Blue  Stain  for  Bacteria  in  Tissues. — 1.  Stain  sections  (paraffin) 
30  minutes  to  24  hours. 

2.  Acetic  acid  (1  to  1,000  of  water),  10  to  20  seconds. 

3.  Rinse  in  absolute  alcohol  20  to  30  seconds. 

4.  Xylol. 

5.  Xylol-balsam. 

With  celloidin  sections  substitute  95  per  cent  alcohol  for  absolute 
(step  3),  then  treat  with  creosote  or  cedar  oil  until  sections  are  clear.  Mount 
in  xylol-balsam. 

Anilin  gentian  violet,  methyl  blue,  methyl  violet,  or  fuchsin  (memo- 
randum 3,  a,  p.  116),  also  carbol-fuchsin  (memorandum  3,  c,  p.  116)  may  be 
used  in  the  same  way. 

Gram's  Method  for  Bacteria  in  Tissues  (Weigert's  modification). — 
1.  Stain  sections  (any  kind)  in  lithium  carmine  2  to  5  minutes. 

Lithium  Carmine  (Orth's): 

Carmine 2 . 5  to  5  grams 

Carbonate  of  lithium,  saturated  aqueous 

solution 100  c.c. 

Thymol a  crystal  or  two 

Filter 

2.  Anilin  gentian  violet,  5  to  20  minutes  (celloidin  sections  should  first 
be  dehydrated  in  95  per  cent  alcohol  and  affixed  to  the  slide  with  ether 
vapor) . 

3.  Rinse  in  normal  saline. 

4.  Gram's  solution  (memorandum  3,  /,  p.  117),  1  to  2  minutes. 

5.  Rinse  in  water. 

6.  Blot    sections    with    filter-paper    to    remove    as   much   water   as 
possible. 

7.  Anilin  oil,  several  changes.     The  oil  dehydrates,  and  at  the  same 
time  decolorizes  the  celloidin. 

8.  Xylol,  several  changes. 

9.  Xylol-balsam. 


Bacteria  115 

C.    Hanging-Drop  Preparations 

1.  A  slide  with  a  concave  center  is  used  (Fig.  40).    With  a  fine-pointed 
brush  paint  a  narrow  strip  of  vaseline  around  the  margin  of  the  concavity. 
The  vaseline  makes  the  cover-glass  stick  to  the  slide  and  also  prevents 
evaporation. 

2.  Place  a  small  drop  of  the  fluid  containing  bacteria  in  the  center  of 
the  cover-glass.    If  the  bacteria  to  be  examined  are  on  a  solid  medium,  the 
"drop"  should  be  made  by  mixing 

a  small  portion  of  the  growth  with    f  :  .  -  .    > ..  .^...J  ^sgsiL?*! 

a  drop  of  bouillon,  normal  saline,  pIQ>  40.— Culture  Slide 

or  serum.    Place  the  cover-glass, 

drop  downward,  over  the  depression  in  the  slide  and  press  it  down  well  into 

the  vaseline. 

3.  Use  only  a  small  opening  in  the  diaphragm  when  examining  the 
bacteria,  in  order  to  get  as  much  contrast  by  refraction  as  possible.    Focus 
first  with  a  medium-power  dry  objective  on  the  edge  of  the  drop,  then 
employ   the   oil  immersion.    Such  unstained   organisms   are  frequently 
difficult  to  find  and  there  is  great  danger  of  breaking  the  cover-glass  with 
the  objective. 

Hanging-drop  preparations  are  used  mainly  in  determining  the  moUHty 
of  bacteria,  or.  in  the  study  of  spore  formation.  For  the  latter  purpose 
the  sEdE5  and  cover-glass  must  be  carefully  sterilized  and  the  sealing  with 
vaseline  complete.  The  preparation  may  then  be  placed  on  a  warm  stage 
or  in  an  incubator  and  examined  from  tune  to  tune. 

MEMORANDA 

1.  The  Main  Points  to  Be  Observed  in  the  Microscopical  Examination 
of  Bacteria  are  as  follows:    (1)  form  of  the  individual,  whether  spherical 
(coccus),  spiral  (spirillum),  or  rodlike  (bacillus)  with  end  square,  pointed, 
or  rounded;    (2)  uniformity  in  size;    (3)  the  arrangements  of  individuals 
whether  single  (micrococci,  etc.),  in  pairs  (e.g.,  diplococci),  in  chains  (e.g., 
streptococci),  groups  of  four  (e.g.,  tetracocci),  cubical  groups  of  eight  or  more 
(sarcinae),  or  small  grapelike  bunches  of  various-sized  cocci  (staphylococci) ; 
(4)  presence  or  absence  of  cell  wall,  gelatinous  capsule,  etc.;    (5)  motility 
in  living  forms  (do  not  confuse  with  Brownian  movement) ;   (6)  reaction  to 
stains;    (7)  presence  of  spores  which  are  recognizable  as  bright,  highly 
refractive  rounded  bodies. 

2.  Material  for  the  Demonstration  of  Bacteria  (coccus,  bacillus,  spiril- 
lum, and  beggiatoa  forms)  will  be  found  in  abundance  in  foul  water,  espe- 
cially when  contaminated  with  sewage.    By  scraping  the  inside  of  the  cheek 
such  forms  as  Leptothrix  may  often  be  found.    Make  a  cover-glass  prepara- 
tion; kill  and  fix  in  the  flame  in  the  ordinary  way;  stain  in  methyl  violet, 


116  Animal  Micrology 

gentian  violet,  or  fuchsin  (basic)  and,  if  desired,  counterstain  lightly  with 
Bismarck  brown;  examine  in  water  or  dehydrate  in  absolute  alcohol,  clear 
in  xylol,  and  mount  in  balsam. 

To  demonstrate  bacteria  in  tissues,  a  mouse  may  be  inoculated  with 
anthrax,  and  paraffin  sections  of  the  spleen  prepared.  Stain  by  the  gentian- 
violet  method. 

3.  Some  of  the  Most  Important  Stains  for  Bacteria  are  as  follows: 

a)  Anilin  water  solution  of  gentian  violet  (Koch-Ehrlich's) . — 

Gentian  violet,  saturated  alcoholic  solution 10  c.c. 

Anilin  water  (see  reagent  30,  p.  218) 100  c.c. 

After  shaking,  the  mixture  should  be  set  aside  for  24  hours  because  of  the 
precipitation  which  takes  place  soon  after  making.  Solutions  of  fuchsin 
(basic)  and  methyl  blue  are  made  in  the  same  way.  These  solutions  begin 
to  decompose  after  about  10  days  and  must  then  be  freshly  prepared.  They 
yield  good  results  with  many  species  of  bacteria.  The  gentian  violet, 
particularly,  is  widely  used  in  connection  with  Gram's  method  (see/). 

6)  Alkaline  methylen  blue  (Loffler's). — 

Methylen  blue,  saturated  alcoholic  solution 30  c.c. 

Caustic  potash,  aqueous  solution  (1 : 10,000) 100  c.c. 

This  stain  keeps  well  and  is  one  of  the  most  widely  used  of  the  general  stains. 
It  is  especially  serviceable  in  staining  the  bacillus  of  diphtheria  or  of  glanders. 

c)  Carbol-fttchsin  (Ziehl-Neelson's). — 

Fuchsin,  saturated  alcoholic  solution 10  c.c. 

Carbolic  acid,  5  per  cent  aqueous  solution 90  c.c. 

This  stain  keeps  well,  stains  powerfully,  and  can  be  used  on  many  forms  of 
bacteria. 

d)  Neisser's  method  for  the  diagnosis  of  diphtheria. — 
Solution  I: 

Methylen  blue  (Griibler's) 1  gram 

Alcohol,  96  per  cent 20  c.c. 

Distilled  water  (add  after  the  methylen  blue  has 

dissolved  in  the  alcohol) 950  c.c. 

Glacial  acetic  acid 50  c.c. 

Solution  II: 

Bismarck  brown 1  gram 

Distilled  water  (should  be  boiling  when  the  Bis- 
marck brown  is  added) 500  c.c. 


Bacteria  117 

Cover-glass  preparations  are  stained  for  from  2  to  3  seconds  in  Solution 
I,  rinsed  in  distilled  water,  placed  in  Solution  II  for  from  3  to  5  seconds, 
rinsed  again  in  water,  and  examined  in  the  ordinary  way.  The  bacteria 
of  virulent  diphtheria  should  appear  as  pale-brown  rods,  some  of  which 
show  at  one  or  both  ends  bluish-black  oval  bodies  of  greater  diameter  than 
the  rod.  Such  dark  bodies  will  not  be  seen  in  the  pseudo-diphtheria  bacilli. 
The  bacilli  must  have  been  grown  for  from  12  to  18  hours  on  Loffler's 
blood-serum  which  is  a  mixture  of  glucose  bouillon  1  part  and  beef-blood 
serum  3  parts.  The  mixture  is  run  into  test-tubes  and  coagulated  at  100°  C. ; 
the  tube  should  be  tilted  to  one  side  to  give  a  slanting  surface  for  culture 
purposes.  The  formula  for  glucose  bouillon  is  as  follows:  dry  glucose, 
10  grams;  Liebig's  extract  of  beef,  3  grams;  peptone,  10  grams;  sodium 
chloride,  5  grams;  water  1,000  c.c. 

e)  Gobbet's  solution  for  demonstrating  tubercle  bacilli. — 

Methylen  blue 1  to  2  grams 

Distilled  water 75  c.c. 

Concentrated  sulphuric  acid 25  c.c. 

The  acid  decolorizes,  while  the  methylen  blue  serves  as  a  contrast  stain. 
The  solution  acts  rapidly.  A  modification  of  the  method  to  be  commended 
is  first  to  stain  the  preparation  with  carbol-fuchsin  (see  c)  by  warming  the 
stain  on  the  slide  until  it  steams,  rinsing  in  water,  and  then  proceeding 
with  the  methylen-blue  solution.  Smegma  and  leprosy  bacilli,  and  the 
treponema  of  syphilis  are  also  stained  by  this  method.  Tubercle  bacilli  are 
also  stained  by  Gram's  method  (see  /).  To  examine  sputum  for  tubercle 
bacilli,  the  sputum  is  carefully  inspected  for  small  yellowish-white  cheesy 
masses  varying  in  size  from  the  diameter  of  a  pin-head  to  that  of  a  small  pea. 
Very  thin  smear  preparations  (see  A,  p.  112)  are  made  from  such  masses. 

/)  Gram's  method. — 

Gram's  solution: 

Iodine  crystals 1  gram 

Iodide  of  potassium 2  grams 

Distilled  water 300  c.c. 

The  preparations  are  first  stained  in  anilin  gentian  violet  (memorandum 
3,  a),  and  then  immersed  in  Gram's  solution  for  from  1  to  2  minutes.  They 
are  then  rinsed  in  alcohol  until  the  violet  color  is  no  longer  visible  to  the  naked 
eye.  To  decolorize  them  sufficiently,  it  may  be  necessary  to  treat  them 
again  with  the  iodine  solution.  Finally  rinse  in  water  and  examine,  or,  if  a 
permanent  preparation  is  desired,  rinse  in  absolute  alcohol,  transfer  to  xylol, 
and  mount  in  balsam.  If  the  preparations  are  from  cultures,  it  should  be 


118 


Animal  Micrology 


borne  in  mind  that  the  method  works  well  only  when  applied  to  bacteria 
from  actively  growing  cultures ;  old  cultures  seldom  yield  satisfactory  results. 


PATHOGENIC  BACTERIA  STAINED 
BY  GRAM'S  METHOD 

Bacillus  aerogenes  capsulatus 
Bacillus  of  anthrax 
Baccilus  diphtheriae 
Bacillus  of  malignant  edema 
Bacillus  of  tetanus 
Bacillus  tuberculosis 
Micrococcus  tetragenus 
Pneumococcus 

Staphylococcus  pyogenes  aureus 
Staphylococcus  pyogenes  albus 
Streptococcus  pyogenes 

Streptococcus  capsulatus 


PATHOGENIC  BACTERIA  DECOLORIZED 
BY  GRAM'S  METHOD 

Bacillus  of  bubonic  plague 
Bacillus  of  chancroid 
Bacillus  coli  communis 
Bacillus  of  dysentery 
Bacillus  of  glanders 
Bacillus  of  influenza 
Bacillus  mucosus  capsulatus 
Bacillus  proteus 
Bacillus  pyocyaneus 
Bacillus  of  typhoid 
Diplococcus  intra  cellularis 

meningitidis 
Gonococcus 
Spirillum  of  Asiatic  cholera 


4.  Staining  Spores  (Abbott's  method). — Prepare  a  cover-glass  smear  in 
the  usual  way.    Apply  the  stain  (e.g.,  methylen  blue)  and  hold  the  cover- 
glass  over  a  flame  until  the  liquid  steams.    Repeat  the  heating  several  times, 
but  do  not  boil  continuously.    Rinse  the  cover-glass  in  water  and  then 
decolorize  the  preparation  in  a  0.3  per  cent  solution  of  hydrochloric  acid 
in  95  per  cent  alcohol,  until  all  color  visible  to  the  naked  eye  has  disappeared. 
Wash  in  water.     If  a  counterstain  is  desired,  stain  for  from  8  to  10  seconds 
in  anilin-fuchsin  solution.    Rinse  in  water  and  mount  in  the  usual  way. 
The  spores  are  stained  blue. 

5.  Staining  Flagella  (Bunge's  modification  of  Loffler's  method). — The 
locomotor  organs  of  motile  bacteria  are  long,  hairlike  prolongations  (1  to 
many)  termed  flagella.     Special  methods  of  staining  are  necessary  for  their 
demonstration. 

Make  thin  cover-glass  smears  of  an  18-hour  culture  which  contains  motile 
forms.  Dry  and  fix  in  the  ordinary  way. 

The  mordant. — 

Ferric  chloride,  aqueous  solution  (1 :20) 25  c.c. 

Alum,  saturated  aqueous  solution 75  c.c. 

Shake  well  and  add 

Fuchsin  (basic),  saturated  aqueous  solution 10  c.c. 

Filter  and  allow  to  stand  for  some  time  before  using.  Treat  the  smear 
for  5  minutes  with  this  preparation,  gently  warming  by  holding  it  high 
above  a  flame.  The  fluid  must  not  boil.  Rinse  in  water,  then  stain  faintly 
with  carbol-fuchsin.  Repeat  the  process  until  a  successful  result  is  obtained. 
Mount  in  the  usual  way. 


CHAPTER  XVI 

SOME  EMBRYOLOGICAL  METHODS;    SECTIONS  AND  "IN 

TOTO"  MOUNTS  OF  FROG  AND  CHICK;  AMPHIBIA; 

FISH;  MAMMALS;  OTHER  FORMS 

THE  FROG 

Frog  eggs  and  tadpoles  are  best  fixed  in  Tellyesnicky's  fluid 
(5,  p.  209).  Eggs  in  early  cleavage  stages  should,  after  such  fixation, 
be  preserved  in  2  per  cent  formalin,  but  later  stages  and  tadpoles  are 
better  preserved  in  70  to  80  per  cent  alcohol.  Before  eggs  can  be 
sectioned,  the  thick  albuminous  coats  which  surround  them  must 
be  removed  (4,  p.  121).  In  addition  to  the  ordinary  cleavage  and 
yolk-plug  stages,  I  find  3,  5,  7,  and  9  mm.  tadpoles,  both  as  whole 
mounts  and  sectioned,  the  most  useful  stages  for  a  course  in  embry- 
ology. Older  stages  are  also  necessary  for  the  study  of  external 
features  of  later  development. 

Allen  (op.  tit.,  p.  122)  finds  that  cleavage  furrows  show  with 
greater  distinctness  if  the  eggs  are  bleached.  The  fixed  and  hardened 
specimens  are  placed  in  ordinary  commercial  hydrogen  peroxide 
for  a  week  or  more  until  the  pigmented  area  is  of  a  light-brown  color. 
If  formalin-hardened  material  is  used,  the  formalin  should  be  washed 
out  before  the  objects  are  placed  in  the  peroxide,  otherwise  the  tissue 
will  be  distorted  by  the  rapid  liberation  of  oxygen.  Tadpoles  may 
be  bleached  white  by  this  method  and  mounted  entire  (see  B,  p.  120). 

A.    Section  Method 

1.  Select  several  7  mm.  tadpoles  which  have  been  fixed  in  Telly  es- 
nicky's  fluid  (5,  p.  209)  and  stain  for  from  12  to  24  hours  in  alum- 
cochineal. 

2.  Run  the  stained  specimens  up  through  the  grades  of  alcohol 
into  absolute  alcohol. 

3.  Transfer  to  absolute  alcohol  and  chloroform,  equal  parts,  for 
an  hour,  then  to  pure  chloroform.     After  an  hour  add  melted  paraffin 

119 


120  Animal  Micrology 

to  the  chloroform  from  time  to  time  until  the  latter  contains  all  the 
paraffin  it  will  hold  in  solution.  Leave  the  objects  in  this  mixture 
for  at  least  24  hours. 

4.  Transfer  to  melted  paraffin  (melting-point  about  48°  C.)  and 
keep  for  2  or  3  hours  at  a  temperature  just  high  enough  to  liquefy 
the  paraffin.     Imbed  after  reading  step  5. 

5.  Prepare  at  least  three  sets  of  sections,  one  set  in  each  of  the 
three  different  planes  of  the  body.     The  sections  should  be  cut  some 
20  or  30  microns  thick.     Read  carefully  the  directions  under  mem- 
orandum 1,  p.  126,  before  imbedding,  so  that  the  sections  will  be 
properly  oriented.     Read  also  caution  under  step  10  on  p.  125. 

6.  Carefully  following  directions  under  memorandum  1,  p.  126, 
mount  the  sections  with  albumen  fixative  and  albuminized  water  as 
usual  (steps  17-21,  pp.  40-41). 

7.  Dissolve  out  the  paraffin  from  the  sections  in  the  usual  way 
after  the  latter  are  thoroughly  dry.     If  not  completely  dry  and  tightly 
stuck  to  the  slide  some  of  the  sections  will  float  off. 

8.  Pass  the  slides  back  into  absolute  alcohol  for  a  few  minutes, 
then  into  fresh  xylol  until  clear.     Add  balsam  and  the  cover-slip. 

B.    Whole  Mounts  (after  Allen,  op.  tit.,  p.  122) 

1.  Select  5  to  7  mm.  tadpoles  which  have  been  fixed  in  Tellyes- 
nicky's  fluid  (5,  p.  209)  and  bleached  in  hydrogen  peroxide  for  one 
week,  or  until  white. 

2.  Stain  for  12  to  24  hours  in  alum-cochineal  diluted  with  distilled 
water  until  the  stain  shows  but  a  faint  tinge  of  color. 

3.  Run  the  tadpoles  up  through  the  increasing  grades  of  alcohol 
into  cedar  oil,  creosote,  or  synthetic  oil  of  wintergreen  and  leave  until 
clear. 

4.  Using  Canada  balsam  or  damar  which  has  been  heated  in  an 
oven  for  some  days  until  it  will  harden  immediately  upon  cooling, 
prepare  several  slides  by  dropping  such  heated  balsam  upon  them 
until  drops  3  or  4  mm.  deep  and  as  wide  as  the  cover-slip  are  formed. 
Let  these  harden.     If  small  bubbles  appear  in  such  drops,  place  the 
slides  in  an  incubator  or  a  paraffin  oven  for  some  hours;  the  bubbles 
rise  to  the  surface  and  may  be  skimmed  off  or  burst  with  a  flame. 


Some  Embryological  Methods  121 

5.  To  mount  a  tadpole  in  such  a  drop,  heat  the  drop  by  holding 
it  inverted  over  a  flame  for  a  moment,  then  with  a  scalpel  previously 
dipped  in  xylol  make  a  groove  in  it  of  suitable  size  and  shape  to  fit 
the  tadpole.     The  sides  of  the  groove  should  press  against  the  object 
in  such  a  way  as  to  hold  it  in  the  desired  position.     Some  tadpoles 
should  be  mounted  to  present  a  dorsal,  others  a  lateral,  aspect. 

6.  When  the  object  is  in  place,  fill  in  the  space  about  it  with  a 
drop  of  soft  balsam  and  gently  but  firmly  press  a  heated  cover-slip 
down  upon  the  surface  of  the  drop.     Guard  against  including  air 
bubbles. 

7.  Examine  the  preparations  after  a  week  or  more  and,  by  pushing 
the  cover-slip  toward  one  side  or  the  other,  adjust  any  of  the  objects 
which  may  have  shifted  from  the  desired  position. 

MEMORANDA  ON  AMPHIBIAN  MATERIAL 

1.  To  Study  Amphibian  Eggs  Entire,  use  a  hand  lens  or  dissecting 
microscope.    Place  the  eggs  on  a  bit  of  absorbent  cotton  under  70  per  cent 
alcohol  in  salt  cellars.    The  eggs  are  fragile;  consequently,  to  manipulate 
them,  use  a  soft-hair  pencil  or  a  current  from  a  pipette.    Use  the  same  egg 
for  surface  view  and  for  sectioning  when  possible. 

2.  Special  Egg  Pipettes  for  handling  delicate  objects  should  be  prepared 
by  breaking  off  the  tip  of  an  ordinary  pipette  to  enlarge  the  orifice.    After 
rounding  up  the  broken  edges  in  a  flame,  cover  the  broken  end  with  a  small 
piece  of  soft-rubber  tubing. 

3.  Amphibian  Eggs  in  General  may  be  fixed  (in  masses  of  15  or  20)  in 
Tellyesnicky's  fluid  (5,  p.  209)  or  in  Worcester's  aceto-formol-sublimate 
mixture  (20,  b,  p.  214).    Chromic  acid  (11,  p.  211)  brings  out  surface  views 
well,  but  the  material  becomes  very  brittle  and  does  not  take  stains  readily. 
If  surface  views  alone  are  desired,  formalin-preserved  material  will  answer. 

4.  To  Remove  the  Gelatinous  Coats  of  Eggs,  roll  them  over  and  over  on 
a  bit  of  blotting  paper.    Either  fresh  or  preserved  eggs  may  be  handled  in 
this  way.    To  prevent  very  soft  eggs  from  drawing  down  and  adhering 
tightly  to  the  blotting  paper,  roll  them  off  on  to  a  paper  of  harder  texture  just 
before  the  last  trace  of  gelatinous  film  has  been  removed. 

Whitman  (American  Naturalist,  XXII,  857)  recommends  putting  the 
fixed  eggs  into  a  10  per  cent  solution  of  sodium  hypochlorite  diluted  with 
5  or  6  volumes  of  water  and  leaving  them  until  they  can  be  shaken  free. 
This  requires  only  a  few  minutes.  Rinse  the  eggs  in  35  per  cent  alcohol. 
It  is  advisable  to  remove  the  albuminous  coats  before  hardening  in 
alcohol. 


122  Animal  Micrology 

Child  (Zeitschrift  fur  wissenschaftliche  Mikroskopie,  XVII  [1900],  205) 
states  that  the  albumen  which  surrounds  many  ova  becomes  transparent 
and  dissolves  if  after  fixation  (in  any  way  except  with  chromic  acid)  the  ova 
are  passed  up  through  the  grades  of  alcohol  to  80  per  cent,  hardened,  and  then 
passed  down  again  through  the  alcohols  into  water  which  has  been  slightly 
modified  with  any  acid  except  chromic. 

5.  Amphibian  Eggs  Are  So  Friable  that  they  are  ordinarily  sectioned  in 
celloidin.    If  they  are  cleared  from  95  per  cent  alcohol  (avoiding  absolute) 
into  cedar  oil  or  oil  of  wintergreen  they  are  less  brittle.    However,  good 
sections  are  obtainable  with  the  Johnson  asphalt-rubber  method  (memo- 
randum 9,  p.  43)  or  by  the  more  tedious  paraffin-celloidin  method  (mem- 
orandum 8,  p.  64) .    Older  embryos  are  readily  sectioned  in  paraffin  according 
to  the  method  already  given  (p.  119). 

6.  Whole  Eggs  May  Readily  Be  Cut  into  Halves  with  a  safety-razor 
blade.    They  are  often  more  serviceable  for  study  along  with  observations 
__________^_^^^^      on  the  external  changes  of  cleavage,  blastula 

formation,  gastrulation,  etc.,  than  the  most 
elaborate  serial  sections. 

7.  The  Germinal  Layers  are  much 
more  distinctly  seen  in  sections  of  young 
embry°s  of  Ambystoma  than  in  those  of 
the  frog. 

8.  Very  Thick  Sections  are  sometimes  useful.    A  7  to  9  mm.  tadpole, 
for  example,  cut  into  three  sagittal  sections  is  excellent  for  studying  the 
general  topography  of  organs.    Also  older  tadpoles  with  the  skin  removed 
from  one  side  before  mounting  are  serviceable. 

9.  Eggs  Mounted  Entire  or  in  Halves,  according  to  Allen's  gelatin 
method  (Kansas  University  Bulletin,  IX,  No.  8  [December,  1914]),  are  very 
successfully  studied  as  opaque  objects.    The  method,  also  useful  for  pre- 
serving free-hand  sections  of  various  kinds,  fine  dissections,  etc.,  is  as  follows: 

a)  Dissolve  thymol  in  distilled  water  with  the  aid  of  heat  until  a  saturated 
solution  is  obtained.     Filter  until  clear. 

b)  Soak  gelatin  in  the  thymol  water  until  it  has  absorbed  all  it  can  hold, 
then  drain  off  the  excess  of  water.    The  gelatin,  now  ready  for  melting,  may 
be  kept  in  corked  test-tubes. 

c)  Prepare  cells  for  mounting  large  objects  by  placing  strips  of  glass  upon 
a  slide  as  indicated  by  the  shading  in  the  diagram  (Fig.  41). 

d)  Liquefy  some  of  the  gelatin  by  placing  a  test-tube  full  into  a  warm 
water-bath,  then  pour  it  into  the  newly  made  cell. 

e)  When  the  gelatin  has  set,  melt  small  areas  in  it  with  a  hot  needle  and 
insert  the  objects  (e.g.,  a  series  of  frog  eggs  in  different  stages  of  segmentation) 
to  be  mounted.    Each  of  these  must  be  held  in  proper  position  with  the  needle 
until  the  gelatin  solidifies. 


Some  Embryological  Methods  123 

/)  Flood  the  cell  with  gelatin  heated  just  enough  to  be  a  liquid  and  place 
a  slightly  warmed  slide  on  top  as  a  cover.  Avoid  air  bubbles,  and  see  that 
there  is  a  complete  film  of  gelatin  between  the  cover-slide  and  the  glass 
strips.  The  cover  should  be  held  firmly  in  place  till  the  gelatin  solidifies. 

g)  With  a  toothpick  or  similar  object  thoroughly  clear  out  every  trace 
of  gelatin  from  the  grooves  formed  by  the  projecting  edges  of  the  slides  and 
the  strips  between  them.  Dry  well. 

h)  Run  some  cement  such  as  gold  size  into  the  groove  and  set  the  prepa- 
ration aside  to  harden.  Add  more  cement  from  tune  to  time  until  the  groove 
is  completely  filled  up.  Keep  the  preparation  out  of  direct  sunlight  or  away 
from  radiators,  as  it  must  not  be  subjected  to  heat. 

If  the  objects  have  previously  been  hardened  in  formalin,  so  much  the 
better,  as  the  formalin  will  gradually  diffuse  out  into  the  gelatin  and  harden  it. 

Hollow-ground  slides  designed  for  use  with  a  hanging  drop  may  be  used 
if  preferred.  It  is  sometimes  desirable  to  solidify  the  gelatin  more  rapidly 
after  the  object  is  mounted,  by  placing  it  on  ice. 

10.  For  Artificial  Fecundation  of  Amphibian  Eggs  see  16,  b,  p.  135. 


THE  CHICK 

A  setting  hen  or  an  artificial  incubator  is  necessary.  In  many 
ways  the  latter  is  more  convenient  as  it  may  be  kept  in  the  laboratory 
and  is  ready  at  all  seasons  of  the  year.  There  are  many  kinds  of 
good  incubators  on  the  market  at  present  which  may  be  had  for  a 
small  sum. 

Whatever  method  of  incubation  is  employed,  the  eggs  must  be 
fresh  and  must  not  have  been  subjected  to  rough  handling.  The 
date  and  hour  at  which  incubation  is  to  begin  should  be  written 
on  the  shell  of  each  egg  in  ink.  If  late  stages  of  development  are 
desired,  the  egg  must  be  turned  every  few  days.  All  products  of 
combustion  from  the  lamp  or  burner  should  be  kept  from  the  eggs 
and  the  supply  of  fresh  air  and  moisture  carefully  maintained.  The 
temperature  should  be  maintained  at  39°  C.  (102°  F.).  Should  it 
rise  above  40°  C.,  embryos  will  be  destroyed. 

Prepare  at  least  5  embryos  as  directed  in  the  practical  exercise, 
2  for  in  toto  preparations  and  3  for  sections. 

1.  Place  an  egg  which  has  been  incubated  for  between  46  and 
54  hours,  while  it  is  yet  warm,  in  a  vessel  which  contains  sufficient 
normal  saline  warmed  to  39°  C.  to  cover  the  egg.  In  the  fowl  the 


124  Animal  Micrology 

embryo  always  makes  its  appearance  as  a  germinal  disk  or  cicatricula, 
as  it  is  termed,  situated  on  one  side  of  the  yolk,  which  is  the  real  egg 
of  the  hen,  the  white  being  simply  a  nutritive  mass  added  in  the 
oviduct.  This  disk  or  blastoderm  in  the  early  stages  of  incubation 
always  turns  uppermost  no  matter  in  what  position  the  egg  may  be 
placed.  Moreover,  it  has  been  found  that  the  embryo  in  nearly 
every  instance  lies  in  such  a  position  that  when  the  blunt  end  of  the 
egg  is  toward  the  left,  the  head  of  the  chick  is  directed  away  from 
the  operator.  This  fact  affords  a  very  reliable  means  of  orienting 
the  embryo,  especially  in  the  very  young  stages  when  the  anterior 
and  posterior  ends  are  not  easily  recognized  by  the  observer.  * 

2.  Break  through  the  shell  at  the  broad  end  over  the  air  chamber 
by  tapping  it  sharply,  and  let  out  the  air,  or  the  broad  end  will  tilt  up. 

3.  Begin  at  the  hole  made  in  the  end  and  with  blunt  forceps 
remove  the  shell  and  shell  membrane  bit  by  bit  from  the  upper  sur- 
face of  the  egg  until  the  embryo  comes  plainly  into  view.     Remove 
with  a  pipette  the  thin  layer  of  albumen  which  lies  above  the  blasto- 
derm. 

4.  With  as  little  agitation  of  the  liquid  in  the  vessel  as  possible 
by  means  of  fine  scissors  cut  rapidly  around  the  blastoderm  well  out- 
side of  the  vascular  area. 

5.  Carefully  float  the  blastoderm  into  a  thin  watch-glass,  keep- 
ing it  as  flat  as  possible.     Shake  it  gently  to  remove  .the  piece  of 
vitelline  membrane  covering  it,  or  any  yolk  which  may  adhere. 
The  aid  of  a  needle  may  be  necessary  to  remove  the  vitelline  covering. 

6.  With  a  pipette  remove  all  excess  of  fluid  from  the  watch- 
glass  but  do  not  let  the  embryo  become  dry.     In  order  to  keep  the 
edges  from  curling  up  and  obscuring  the  embryo,   touch  small, 
rectangular  pieces  of  dry  filter-paper  to  the  blastoderm  around  its 
periphery  (Miller's  method:   Anatomical  Record,  V,  No.  8  [August, 
1911]),  and  when  they  adhere  use  them  for  spreading  the  tissues 
flat.     Stick  the  free  ends  of  the  paper  strips  to  the  bottom  of  the 
watch-glass  to  hold  the  membranes  in  this  spread  condition. 

7.  Carefully  add  picro-sulphuric  fixer  (reagent  26,  p.  217)  until 
the  embryo  is  completely  immersed.     The  fluid  should  be  allowed 
to  act  for  from  2  to  3  hours. 


Some  Embryological  Methods  125 

The  paper  strips  may  be  left  in  place  through  subsequent  treat- 
ment to  95  per.  cent  alcohol  for  embryos  which  are  to  be  sectioned,  or 
to  xylol  for  those  which  are  to  be  mounted  whole. 

NOTE. — Some  prefer  to  fix  the  embryo  before  removing  it  from  the  egg. 
After  some  of  the  albumen  is  drawn  off,  the  fixing  agent  is  squirted  on  to  the 
blastoderm.  As  soon  as  it  is  opaque  the  latter  is  then  removed  to  a  vessel  con- 
taining the  fixing  agent.  Andrews  (Zeitschrift  fur  wissenschaftliche  Mikroskopie, 
XXI  [1904],  177)  injects  picro-sulphuric  acid  (1)  between  the  vitelline  membrane 
and  the  blastoderm  and  (2)  between  the  blastoderm  and  the  yolk  by  means  of 
a  pipette  which  has  a  fine  upcurved  point.  The  blastoderm  may  then  be  readily 
freed  from  the  yolk.  This  operation  should  be  performed  before  the  egg  has 
been  subjected  to  the  action  of  any  reagents. 
% 

8.  Wash  in  repeated  changes  of  70  per  cent  alcohol.     Pass  down 
through  50  and  35  per  cent  alcohol  to  water.     Stain  in  alum-cochineal 
for  24  hours  (Conklin's  hematoxylin  may  be  used  if  preferred). 

9.  Wash  the  object  in  water  and  transfer  it  through  35  and  50 
per  cent  alcohol,  leaving  it  30  minutes  in  each.     Decolorize  the 
embryo  slightly  in  weak  acid  alcohol,  then  wash  in  70  per  cent  alcohol, 
and  leave  it  -there  until  ready  to  proceed. 

10.  Transfer  the  object  through  95  per  cent  (1  hour),  absolute 
alcohol  (2  hours)  to  xylol,  where  it  should  remain  about  2  hours  or 
until  it  ceases  to  appear  opaque. 

Mount  two  embryos  entire,  one  with  the  ventral,  the  other 
with  the  dorsal,  side  uppermost.  Put  bits  of  broken  cover-glass 
or  threads  of  glass  under  the  edges  of  the  cover  to  avoid  crushing 
them. 

The  three  remaining  embryos  are  to  be  so  sectioned  (steps  11  ff.) 
that  the  student  will  have  a  complete  series  of  sections  in  each  of  the 
three  different  planes  of  the  body  with  reference  to  the  axis  of  the 
spinal  cord:  viz.,  transverse,  sagittal,  and  frontal.  Read  carefully 
memorandum  1  on  orienting  serial  sections. 

CAUTION. — Before  sectioning  any  embryo  always  make  an  outline 
drawing  of  the  entire  embryo;  then  rule  lines  across  the  drawing  parallel 
to  the  plane  of  section.  Unless  this  is  done  great  difficulty  will  be  experi- 
enced frequently  in  understanding  the  sections. 

11.  Infiltrate  the  embryo  with  paraffin  in  the  usual  manner  by 
leaving  it  in  melted  paraffin  for  2  or  3  hours.     A  paraffin  melting  at 


126  Animal  Micrology 

about  48°  C.  should  be  employed  and  sections  should  be  cut  20  to 
30  microns  thick. 

12.  Imbed  and  cut  in  the  usual  way  (chap.  v).     Mount  the  entire 

series. 

MEMORANDA 

1.  Directions  for  Orienting  Serial  Sections. — a)  In  mounting  transverse 
sections  (sections  across  the  main  axis  of  the  object),  the  sections  beginning 
at  the  anterior  end  of  the  object  are  laid  on  the  slide  in  the  same  sequence 
as  the  reading  on  the  page  of  a  book.     In  order  to  have  right  and  left  sides 
and  dorsal  and  ventral  surfaces  in  proper  relation  to  the  observer,  mount 
the  object  in  such  a  way  that,  in  cutting,  the  knife  will  enter  it  on  the  left 
side  and  at  the  anterior  end.    Leave  room  at  one  end  of  the  slide  (see  p.  £9) 
for  a  label  and  also  a  small  margin  at  the  opposite  side. 

6)  To  get  proper  orientation  of  frontal  sections  (sections  lengthwise  of 
the  object  in  a  plane  including  right  and  left  sides),  arrange  the  object  so 
that  the  knife  will  enter  it  on  the  right  side  and  slice  off  the  dorsal  surface 
first.  Mount  sections,  with  their  posterior  ends  toward  the  upper  edge  of 
the  slide,  placing  the  first  section  of  the  series  to  the  left  end  of  the  upper 
row.  This  throws  left  and  right,  dorsal  and  ventral,  into  their  proper  posi- 
tion as  viewed  through  the  compound  microscope,  and  the  observer  looks 
from  the  dorsal  toward  the  ventral  aspect  of  the  object. 

c)  To  mount  sagittal  sections  (sections  lengthwise  of  the  object  in  a  plane 
including  ventral  and  dorsal  sides),  arrange  the  object  in  such  a  position  that 
the  knife  enters  the  ventral  surface  and  slices  off  the  right  side  first.  Mount 
with  the  posterior  end  toward  the  upper  edge  of  the  slide,  placing  the  first 
section  of  the  series  at  the  left  end  of  the  upper  row.  Through  the  com- 
pound microscope  the  observer  views  the  object  from  the  right  toward  the 
left.  The  head  will  appear  to  be  toward  the  upper  end  of  the  slide,  the  dorsal 
surface  toward  the  left. 

It  is  frequently  advantageous  to  have  the  imbedding-mass  trimmed 
unsymmetrically  by  leaving  the  edge  which  first  comes  in  contact  with  the 
knife  longer  than  the  opposite  edge.  One  may  thus  readily  discover  if  a 
section  or  part  of  a  series  has  been  accidentally  turned  over. 

2.  Orientation  of  Objects  in  the  Imbedding-Mass  so  that  sections  can 
be  cut  accurately  in  definite  planes  is  frequently  difficult  to  accomplish.    The 
following  methods  are  useful  in  many  instances: 

I.  For  paraffin  sections. — With  a  soft  pencil  rule  the  strip  of  paper  which 
is  to  be  used  for  making  the  imbedding-box  into  small  squares  or  rectangles. 
After  imbedding,  upon  removal  of  the  paper  a  copy  of  the  pencil  marks  will 
be  found  upon  the  block  of  paraffin.  If  the  object  has  been  arranged  in 
the  melted  paraffin  with  reference  to  these  lines,  it  is  easy  so  to  arrange 
the  block  in  the  microtome  as  to  cut  the  object  along  any  desired  plane. 


Some  Enibryological  Methods  127 

It  is  frequently  an  aid  to  orientation  by  this  method  to  have  one  of  the  central 
ruled  lines  broader  than  the  others,  or  double. 

Small  objects  which  cannot  conveniently  be  oriented  in  melted  paraffin 
may  be  properly  oriented  and  fixed  to  a  small  strip  of  paper  ruled  as  above, 
before  they  are  placed  in  the  paraffin  bath,  by  a  mixture  of  clove  oil  and 
collodion  of  about  the  consistency  of  thick  molasses,  as  in  Patton's  method 
(Zeitschrift  fur  wissenschaftliche  Mikroskopie,  XI  [1894],  13).  One  or  a 
number  of  small  objects  which  have  previously  been  cleared  in  oil  of  bergamot 
or  cloves  are  mounted  in  small  separate  droplets  of  the  reagent  and  oriented 
under  a  dissecting  lens  with  reference  to  the  ruled  lines.  The  paper  is  then 
placed  in  turpentine  which  washes  out  the  clove  oil  and  fixes  the  object  in 
place.  The  paper  with  objects  attached  is  then  passed  through  melted 
paraffin  and  imbedded  in  the  ordinary  way.  Upon  removal  of  the  paper 
from  the  hardened  block  a  sufficient  number  of  pencil  marks  remain  tc  be 
used  as  a  guide  in  sectioning.  Instead  of  pencil  marks  Pattern  employed 
ribbed  paper. 

II.  For  celloidin  sections. — 

Eycleshymer's  Methods. — a)  For  imbedding,  metal  boxes  made  of 
two  L's  (Fig.  30)  are  used.  The  L's  are  held  together  by  overlapping  strips. 
The  ends  and  sides  of  the  box  are  perforated  at  regular  intervals  by  small 
holes  which  have  been  drilled  opposite  one  another  in  such  a  way  that 
threads  drawn  through  them  are  parallel.  Threads  of  silk  are  run 
through  the  holes  from  side  to  side,  drawn  taut,  and  cemented  to  the  out- 
side of  the  box  with  a  drop  of  celloidin.  Each  piece  of  thread  should  have  an 
end  two  or  three  inches  long  hanging  outside  the  box.  A  piece  of  heavy 
blotting  paper  is  used  as  a  bottom  for  the  box.  The  object  is  oriented  on 
the  parallel  threads  and  the  imbedding-mass  poured  in  and  hardened.  The 
loose  ends  of  the  threads  are  then  soaked  in  a  solution  of  thin  celloidin  which 
contains  lamp-black,  the  celloidin  drops  holding  the  threads  taut  are  dissolved 
by  a  drop  of  ether-alcohol,  and  the  blackened  ends  are  drawn  through  the 
block  of  celloidin.  The  lamp-black  leaves  distinct  black  lines  through  the 
mass  which  will  serve  for  properly  orienting  the  celloidin  block  on  the  micro- 
tome. 

This  method  is  valuable  also  in  reconstructions  from  sections  (see  chap, 
xviii).  In  such  work  it  is  very  desirable  to  establish  "reconstruction  points" 
to  guide  in  fitting  the  wax  plates  together  properly.  The  black  rings  of 
lamp-black  left  in  the  sections  answer  admirably  for  this  purpose. 

6)  For  small  objects  in  which  reconstruction  points  are  not  required 
Eycleshymer  uses  fine  insect  pins  from  which  the  heads  have  been  clipped 
and  the  headless  ends  loosely  inserted  in  handles.  The  objects  are  mounted 
on  the  points  of  the  pins  and  oriented  in  the  desired  position.  Each  pin  is 
then  removed  from  its  handle,  and  the  free  end  is  inserted  from  below  into 
a  small  perforation  which  has  been  made  by  passing  a  somewhat  larger  pin 


128  Animal  Micrology 

lengthwise  through  a  cork.  A  number  of  pins  may  be  mounted  on  the  same 
cork.  To  prevent  the  objects  from  becoming  dry,  the  cork  must  frequently 
be  inserted  into  the  mouth  of  a  vial  full  of  alcohol  in  such  a  way  that  the 
objects  are  immersed.  If  desired,  the  objects  may  be  sketched  in  situ  under 
alcohol  by  weighting  the  cork  with  lead  and  placing  it  in  a  beaker  of  alcohol. 
To  pass  the  objects  through  the  various  grades  of  alcohol,  etc.,  simply  trans- 
fer the  cork  bearing  them  to  successive  vials  of  proper  size  containing  the 
different  fluids.  For  imbedding  in  celloidin-use  the  method  given  on  p.  59, 
steps  2  ff .  When  the  celloidin  mass  has  hardened,  the  paper  is  removed  and 
the  pins  are  drawn  out  through  the  cork,  thus  leaving  the  objects  in  place 
ready  for  sectioning. 

3.  In  Measuring  the  Length  of  Embryos  some  embryologists  (e.g.,  Mjnot) 
measure  the  greatest  length  of  the  embryo  along  a  straight  line  (limbs  not 
included)  when  the  embryo  is  in  its  normal  attitude;  consequently  in  some 
early  stage*  where  the  embryo  is  greatly  flexed  the  neck-bend  would  be  the 
point  to  which  to  measure  instead  of  the  tip  of  the  head,  because  it  is  the  most 
anterior  region;  in  stages  where  the  embryo  is  straight,  the  head  would  be 
included.    Other  embryologists  (e.g.,  His  and  German  authors  in  general) 
make  use  exclusively  of  the  so-called  "neck-length";   that  is,  the  distance 
in  a  straight  line  between  the  neck-bend  and  the  caudal-bend.     Still  others, 
in  the  case  of  human  embryos,  use  the  so-called  "sitting  height"  and  "stand- 
ing height." 

4.  For  the  Embryology  of  Teleosts  the  following  are  the  most  useful 
mounted  stages: 

I.  Whole  mounts.— The  2-,  4-,  8-,  16-,  32-,  and  64-cell  stages  (only  thex 
blastodisk  segments);    early  periblast;    late  periblast;    early  germ-ring; 
embryonic  shield;  various  stages  of  early  embryos,  such  as  embryos  of  45, 
50,  and  60  hours. 

II.  Sections  (paraffin). — Of  4,  16,  and  32  cells  (vertical  sections  parallel 
to  the  first  plane  of  cleavage) ;  late  cleavage  (vertical  sections) ;  early,  mid, 
and  late  periblast  (vertical  sections);    transverse  and  sagittal  sections  of 
early  germ-ring,  embryonic  shield,  early  embryo,  late  germ-ring,  and  closing 
of  blastopore,  respectively. 

All  stages  may  be  fixed  in  picro-acetic  (reagent  24,  p.  216)  or  Bouin's 
fluid  for  30  to  40  minutes.  The  eggs  are  finally  preserved  in  83  per  cent 
alcohol.  Child  finds  that  fixation  for  about  a  minute  in  10  per  cent  acetic 
acid  saturated  with  corrosive  sublimate,  followed  by  10  per  cent  formalin, 
gives  good  results  without  the  yolk  becoming  hard.  The  ova  of  the  Sal- 
monidae  must  be  removed  (after  fixing  and  hardening)  from  their  envelopes 
before  the  embryo  can  be  studied. 

Before  the  preserved  material  can  be  mounted  in  toto  or  sectioned,  the 
essential  part  (the  blastoderm)  must  ordinarily  be  dissected  off  under  a 
dissecting  lens  by  means  of  sharp  needles.  If  the  blastoderms  are  to  be 


Some  Embryological  Methods  129 

mounted  entire  they  may  be  passed  down  through  the  alcohol  (see  Walton's 
device,  memorandum  4,  p.  30),  stained  in  Conklin's  hematoxylin  (reagent  51, 
p.  225),  then  dehydrated  and  mounted  in  the  usual  way.  To  avoid  crushing 
the  objects,  the  cover-glass  should  be  supported  by  means  of  bits  of  broken 
cover  or  glass  threads.  Material  which  is  to  be  sectioned  may  be  stained 
in  toto  or  the  sections  may  be  stained  on  the  slide.  In  the  latter  event,  to 
facilitate  orientation,  it  is  necessary  to  tinge  the  blastoderms  slightly  with 
Bordeaux  red  or  some  other  cytoplasmic  stain  unless  the  fixing  reagent  has 
already  done  so.  For  the  same  reason  it  is  best  to  imbed  the  material  in  a 
watch-glass,  arranging  it  near  the  bottom  of  the  paraffin-mass  so  that  one 
can  see  with  a  microscope  how  to  shape  the  paraffin  block  in  order  to  cut 
sections  in  the  proper  plane.  The  immersion  in  the  melted  paraffin  should 
not  be  longer  than  5  or  10  minutes.  The  paraffin  is  best  hardened  under  95 
per  cent  alcohol.  The  sections  may  be  stained  by  any  of  the  hematoxylin 
methods;  iron-hematoxylin  (p.  51)  yields  excellent  results. 

5.  To  Study  Living  Eggs  of  Teleosts,  a  thin,  flexible  piece  of  sheet  cellu- 
loid or  mica  should  be  used  instead  of  a  cover-glass.    The  egg  must  be  rotated 
from  time  to  time,  and  this  is  easily  accomplished  with  such  a  flexible  cover. 

6.  For  Artificial  Fecundation  of  Teleost  Eggs,  see  p.  135. 

7.  Tilting  the  Microscope  into  a  Horizontal  Position  and  examining  the 
egg  in  its  normal  medium  by  direct  light  is  an  excellent  method  of  studying 
blastodisk  formation  in  such  forms  as  Ctenolabrus,  for  instance.    Inasmuch 
as  the  blastodisk  forms  on  the  lower  side  of  the  egg,  it  appears  to  be  on  top 
when  viewed  through  the  compound  microscope. 

8.  To  Preserve  Teleost  Eggs  in  Convenient  Form  for  Demonstrating 
discoidal  cleavage,  embryonic  shield,  germ-ring,  etc.,  Smith  (Transactions 
of  the  American  Microscopical  Society,  XXXIII,  No.  1  [January,  1914])  seals 
pieces  of  f-inch  glass  tubing  at  one  end  by  holding  in  a  flame.    A  series  of  eggs 
fixed  in  corrosive-acetic  mixture  and  preserved  in  formalin  is  placed  in  each 
tube  and  the  opening  plugged  with  cotton.    The  tube  may  be  held  in  the  hand 
and  examined  with  a  lens  or  dropped  into  a  watch-glass  filled  with  water 
and  examined  under  a  lens  or  binocular  microscope. 

9.  For  the  Average  Course  in  Embryology  of  the  Chick  the  following 
mounted  stages  are  the  most  useful: 

I.  Mounted  in  toto. — 
Approximately, 

48    hours  viewed  from  above  and  below 

oa          ti  ((  it  a  «  « 

30 

24 

18 

12 

64-72  " 

96  hours  (studied  in  alcohol  under  the  dissecting  microscope) 


130  Animal  Micrology 

II.  Sections. — 

48  hours,  transverse,  sagittal,  and  frontal 
36      "  "  " 

30      "  "  " 

24      «  tt  tt 

18      " 

10      "  " 

70       "  t(  (t          tl          ft 

96      "  "  " 

The  number  of  embryos  needed  for  the  above-mentioned  preparations 
is  as  follows : 

5  embryos  of  48  hours  (27-29  somites) 
4        "         "  36      "      (15-18        "      ) 

3  "         "  30      "      (10-14        "      ) 

4  "         "  24      "      (  4-6         "      ) 

2  "  "  18      " 
1  tl  t(  12       " 
1  "  "  60      " 

3  "  "  64-72  hours  (cervical  flexure  formed) 
3  "  96  hours 

10.  To  Mark  Anterior  and  Posterior  Ends  of  Young  Chick  Embryos  in 

blastoderms  which  still  have  a  homogeneous  aspect,  Duval's  osmic-acid 
method  is  very  useful.  With  a  strip  of  paper  5  mm.  wide  by  50  mm.  long 
a  triangular  bottomless  box  with  narrow  base  is  constructed.  This  is  placed 
on  the  yolk  inclosing  the  blastoderm  in  such  a  position  that  the  base  of  the 
triangle  corresponds  to  what  will  be  the  anterior  region  of  the  embryo  (for 
orientation  of  embryo  in  the  egg,  see  step  1  of  the  practical  exercise).  Press 
the  box  down  against  the  yolk  and  fill  it  with  a  0 . 3  per  cent  aqueous  solution 
of  osmic  acid.  In  a  short  time  the  preparation  begins  to  darken  and  the 
osmic  acid  should  be  removed.  The  blastoderm  may  then  be  removed  in  the 
ordinary  manner  and  fixed  as  desired  (Duval  used  chromic  acid  for  fixing). 
However,  it  is  very  difficult  to  separate  the  blastoderm  from  the  egg  during 
the  first  24  hours  of  incubation,  and  it  is  advisable,  therefore,  to  fix  and  harden 
both  together  and  to  remove  the  blastoderm  later  (see  note  under  7,  p.  125). 
The  blackened  area  affords  a  convenient  means  of  orienting  the  preparation 
for  sectioning. 

11.  For  the  Stages  of  Maturation,  Fertilization,  and  Segmentation  in 
Mammals  white  mice  will  prove  most  useful  because  these  processes  are  better 
known  in  them  than  in  other  mammals;   furthermore,  an  abundance  of 
material  may  be  procured.    The  ovum,  however,  is  extremely  small,  measur- 
ing only  about  59  microns  in  diameter.    It  is  surrounded  by  a  very  thin 
zona  pellucida  (1.2  microns).    Long  and  Mark  find  a  modified  Zenker's 
fluid  the  most  satisfactory  for  fixation.    They  make  up  two  solutions:  (1)  a 


Some  Embryological  Methods  131 

4  per  cent  aqueous  solution  of  potassium  bichromate;  (2)  a  4  per  cent 
aqueous  solution  of  corrosive  sublimate  and  20  per  cent  acetic  acid*  The 
two  solutions  are  mixed  in  equal  proportions  when  needed  for  fixing. 

Female  mice  are  in  heat  soon  after  parturition.  They  tend  to  ovulate 
every  21  days  during  the  spring  months.  The  maturation  process  requires 
from  4  to  15  hours  and  usually  occurs  between  13  and  29  hours  after  parturi- 
tion. The  second  polar  spindle  usually  forms  immediately  before  ovulation, 
but  the  second  polar  body  may  not  be  extruded  unless  the  egg  is  fertilized. 
Ovulation  occurs  between  14  and  29  hours  after  parturition.  The  eggs  are 
easily  visible  at  first  in  a  fold  of  the  oviduct  near  the  ovary.  Insemination 
is  most  successful  when  it  occurs  between  18  and  30  hours  after  parturition. 
The  spermatozoa  reach  the  eggs  in  the  upper  end  of  the  Fallopian  tube  in 
from  4  to  7  hours. 

For  details  and  bibliography  see  (1)  Kirkham,  Biological  Bulletin,  XII, 
No.  4  (1907);  also  Transactions  Connecticut  Academy  of  Arts  and  Sciences, 
XIII;  (2)  Long  and  Mark,  "The  Maturation  of  the  Egg  of  the  Mouse," 
Publications  of  the  Carnegie  Institute  of  Washington,  D.C.,  1911;  also,  "The 
Living  Eggs  of  Rats  and  Mice  with  a  Description  of  Apparatus  for  Obtaining 
and  Observing  Them,"  University  of  California  Publications  in  Zoology, 
IX,  No.  3  (Feb.  23,  1912);  (3)  Daniels,  "Mice,  Their  Breeding  and  Rearing 
for  Scientific  Purposes,"  American  Naturalist,  October,  1912;  (4)  see  also 
Danforth,  Anatomical  Record,  X,  No.  4  (February,  1916),  for  some  very 
practical  suggestions  regarding  use  in  classes. 

The  phenomena  of  maturation  and  fertilization  in  the  albino  rat  are  de- 
scribed in  a  paper  by  Sobotta  and  Burckhardt  in  Anatomische  Hefte,  XLII, 
1911  (summarized  in  Huber's  paper  cited  on  p.  133).  See  also  Kirkham 
and  Burr,  "The  Breeding  Habits,  Maturation  of  the  Eggs  and  Ovulation 
of  the  Albino  Rat,"  American  Journal  of  Anatomy,  XV,  1913. 

12.  For  Early  Stages  of  the  Mammalian  Embryo  rabbits  are  commonly 
employed  because  they  breed  readily,  especially  in  the  spring  of  the  year, 
and  the  observer  can  note  the  exact  time  when  the  female  is  covered  if  she 
has  been  kept  separate  from  the  buck  until  she  comes  into  heat.  The  period 
of  gestation  is  30  days  and  impregnation  takes  place  again  immediately 
after  littering.  The  two  uteri  of  the  rabbit  diverge  as  two  anterior  horns 
from  the  single  median  vagina  and  each  terminates  in  front  in  a  narrow,  coiled 
tube,  the  oviduct  or  Fallopian  tube.  To  obtain  the  early  stages  the  abdomen 
is  slit  open  from  pubis  to  sternum,  the  intestinal  tract  is  cut  away  or  pushed 
to  one  side,  and  each  uterus  and  oviduct  carefully  removed  and  stretched  out 
along  a  glass  plate.  The  segmenting  ova  are  found  in  the  oviduct  up  to 
nearly  70  hours  from  the  time  of  copulation.  After  that  period  of  tune  they 
must  be  looked  for  in  the  uterus.  Fecundation  takes  place  about  9  hours 
after  coition.  While  in  the  oviduct,  with  the  aid  of  a  lens  they  may  some- 
times be  seen  through  its  walls.  A  segmenting  ovum  once  located,  a  transverse 


132  Animal  Micrology 

cut  is  made  to  one  side  of  it  through  the  wall  of  the  oviduct,  and  the 
ovum,  which  is  very  small,  is  gently  squeezed  out  by  compressing  the  oviduct 
behind  it.  With  a  spear-headed  needle  or  the  point  of  a  scalpel  the  ovum 
is  conveyed  to  the  fixing  fluid.  In  case  segmenting  ova  are  not  visible  from 
the  exterior  of  the  oviduct,  the  latter  must  be  slit  open  carefully  with  a  pair 
of  fine-pointed  scissors,  and  the  eggs  sought  for  by  means  of  a  lens.  In  case 
no  red  corpora  lutea  are  visible  on  the  surface  of  the  ovary,  indicating  a 
recent  discharge  of  ova  from  the  Graafian  follicles,  further  search  is  useless. 

Rabbit  ova  of  18  hours  show  4  blastomeres;  36  to  48  hours,  advanced 
segmentation;  72  hours  (about  0.6mm.  in  diameter;  in  anterior  end  of 
uterus)  show  the  fully  segmented  ovum — an  outer  layer  of  clear,  cubical 
cells,  an  inner  mass  of  irregular  granular  cells;  72  to  90  hours  show  enlarged 
blastodermic  vesicle  and  establishment  of  embryonic  area;  5th  and  6th 
days  (0.8  to  4mm.)  show  germinal  layers;  7th  day,  primitive  streak;  8th 
day,  medullary  folds. 

The  earlier  stages  (up  to  70  hours)  may  be  fixed  for  from  5  to  8  minutes 
in  a  0 . 3  per  cent  aqueous  solution  of  osmic  acid,  stained  in  picro-carmine, 
and  transferred  to  a  mixture  of  glycerin  and  water,  equal  parts.  They  should 
remain  in  this  fluid  for  a  week  under  a  bell-jar  so  that  the  water  gradually 
evaporates.  The  object  may  then  be  mounted  in  formic-glycerin  (formic 
acid  1  part,  glycerin  99  parts).  To  avoid  pressure  of  the  cover-glass,  the 
object  should  be  mounted  in  a  cell  or  between  two  slips  of  paper  or  pieces  of 
cover-glass.  If  the  preparation  is  to  be  permanent  the  cover-glass  should 
be  sealed  (see  p.  95). 

To  render  the  cell  outlines  distinct,  stages  of  from  70  to  80  hours  are 
best  treated,  after  rinsing  in  distilled  water,  with  a  1  per  cent  aqueous 
solution  of  silver  nitrate  for  3  minutes  and  then  exposed  to  light  in  a  dish 
of  distilled  water  until  they  become  brown.  They  are  then  treated  with 
water  and  glycerin  and  mounted  in  formic-glycerin  as  in  the  case  of  younger 
stages. 

For  sections  the  embryos  should  be  placed  in  Bouin's  or  Zenker's  fluid 
for  one  or  two  hours,  then  washed  in  the  customary  way  for  these  methods, 
stained  in  alum-cochineal,  and  sectioned  in  paraffin. 

In  opening  the  uterus,  the  incision  should  always  be  made  along  the 
middle  of  the  free  side,  opposite  the  insertion  of  the  peritoneal  fold,  because 
this  line  of  insertion  marks  the  region  of  attachment  of  the  embryo  within 
the  oviduct.  By  the  7th  or  8th  day  the  developing  ova  have  taken  up 
positions  at  intervals  along  the  inner  walls  of  the  uterus  and  have  become 
so  firmly  attached  to  the  mucous  membrane  that  they  can  no  longer  be 
detached  unmutilated.  For  further  particulars  regarding  the  embryology 
of  the  rabbit,  the  reader  is  referred  to  E.  Van  Beneden  and  Charles  Julin's 
"Recherches  sur  la  formation  des  annexes  foetalis  chez  les  mammiferes," 
Archives  de  Biologie,  V  (1884),  378.  See  also  Assheton,  "A  Reinvestigation 


Some  Embryological  Methods  133 

into  the  Early  Stages  of  the  Development  of  the  Rabbit,"  Quarterly  Journal 
Microscopical  Science,  XXXVII  (1895). 

With  the  aid  of  Huber's  paper  (Journal  of  Morphology,  XXVI,  No.  2 
[1915];  also  Memoirs  of  the  Wistar  Institute  of  Anatomy  and  Biology,  No.  5), 
which  covers  the  development  of  the  albino  rat  from  the  pronuclear  stage 
to  the  end  of  the  9th  day,  it  is  now  feasible  to  use  the  rat  for  early  embryonic 
stages.  Huber  found  Carney's  fixing  fluid  (reagent  2,  b,  p.  207)  the  most 
satisfactory.  He  fixed  tissues  for  several  hours,  washed  them  in  several 
changes  of  absolute  alcohol,  and  stored  them  in  the  latter.  He  found  that 
sectioning  ovary,  oviduct,  and  uterus  en  masse  was  more  satisfactory  than 
isolating  and  sectioning  the  separate  ova,  although  he  used  both  methods. 
For  staining  he  employed  mainly  hemalum,  followed  by  Congo  red.  His 
methods  are  described  at  some  length  in  the  paper. 

13.  For  Older  Stages  of  the  Mammalian  Embryo  pig  embryos  are  com- 
monly employed.  They  may  often  be  procured  in  large  numbers  and 
with  little  trouble  at  the  larger  pork-packing  establishments.  The  most 
valuable  stage  for  study  is  an  embryo  of  from  10  to  13  mm.  in  length.  In 
most  laboratories  it  is  customary  to  make  a  detailed  study  of  an  embryo  of 
about  this  stage  and  then  a  more  general  survey  of  both  smaller  and  larger 
sizes. 

Early  stages  are  much  more  difficult  to  obtain  than  advanced  stages. 
Embryos  of  6  mm.  length  and  over  may  usually  be  readily  located  by  the 
enlargements  which  they  cause  in  the  uterine  walls.  The  uterus  should  be 
handled  carefully  and  opened  as  soon  as  possible.  The  embryo  is  best 
removed  by  means  of  fine  forceps  and  a  horn  spoon.  It  is  very  delicate 
and  should  not  be  handled  roughly.  The  chances  are  that  in  removing  the 
embryo  the  membranes  will  be  ruptured  and  the  amniotic  and  allantoic 
fluids  will  escape.  Larger  embryos  should  have  the  body  cavity  punctured 
to  admit  the  fixing  fluid. 

Submerge  the  embryo  without  removing  the  membranes  in  a  bounti- 
ful supply  of  Kleinenberg's  picro-sulphuric  acid  (reagent  26,  p.  217),  moving 
it  about  gently  to  rinse  off  any  coagulum  that  may  form  on  the  surface. 
Lavdowsky's  fluid  (19,  p.  214)  is  also  a  good  fixing  agent  for  pig  embryos  and 
is  to  be  preferred  for  the  older  ones. 

Leave  embryos  of  6  to  9  mm.  2§  hours;  12  to  15  mm.,  4  hours;  20  to 
25  mm.,  6  to  8  hours. 

For  washing  and  subsequent  treatment  see  reagent  26,  p.  217.  Embryos 
may  be  stained  in  toto  in  alum-cochineal  or  borax-carmine. 

For  studying  the  uterus,  placentation  (diffuse  in  the  pig),  and  embryonic 
membranes  in  place,  formalin-hardened  material  may  be  used  after  first 
thoroughly  washing  it  in  water. 

For  gross  dissection  of  embryos  the  specimen  should  be  studied  in  alcohol 
under  the  dissecting  microscope. 


134  Animal  Micrology 

Because  of  the  asymmetry  of  young  embryos  it  is  impossible  to  secure 
strictly  transverse,  sagittal,  and  frontal  sections.  Minot  recommends,  there- 
fore, that  for  practical  purposes  the  plane  of  section  be  taken  with  regard 
to  the  head  alone  irrespective  of  how  it  may  cut  the  other  parts  of  the  body, 
and  suggests  the  floor  of  the  fourth  ventricle  of  the  brain  as  the  guide  for 
orientation.  In  his  Laboratory  Text-Book  of  Embryology  he  especially 
recommends  that  each  student  prepare  sections  of  the  following  stages  of 
pig  embryos:  9mm.,  transverse  and  sagittal,  frontal  of  the  head;  6mm., 
transverse,  frontal  of  the  head;  17  mm.,  transverse  and  sagittal,  frontal 
of  the  head;  20  mm.,  transverse  and  sagittal,  frontal  of  the  head;  24  mm., 
frontal  of  the  head. 

14.  Human  Embryos  of  all  ages  are  very  valuable  material  for  scientific 
purposes.    Physicians  and  surgeons  are  urged  to  preserve  such  material 
properly  and  turn  it  over  to  some  competent  embryologist.    Very  young 
human  embryos  are  exceedingly  desirable.    Fill  the  containing  vessel  com- 
pletely with  fluid  in  order  to  avoid  shaking. 

An  excellent  fixing  reagent,  the  ingredients  of  which  a  physician  can 
usually  readily  procure,  is  Lavdowsky's  mixture  (reagent  19,  p.  214).  The 
embryo  should  remain  in  this  fluid  from  12  to  48  hours,  according  to  size,  and 
then  be  preserved  in  80  per  cent  alcohol  (or  commercial  alcohol  to  which 
has  been  added  about  one-fifth  its  volume  of  distilled  water).  Use  a  wide- 
mouthed  bottle  with  tightly  fitting  stopper. 

Zenker's  fluid  (reagent  6,  p.  209)  is  better  for  larger-sized  embryos. 
Material  should  be  left  in  it  from  18  hours  to  several  days.  For  washing 
and  preserving  follow  the  directions  given  under  the  description  of  the  fluid. 
For  fetuses  use  a  fruit- jar  of  such  a  size  that  the  embryo  can  be  kept  in  about 
10  times  its  volume  of  fluid. 

In  case  the  above-mentioned  fluids  are  not  available,  the  material  may  be 
placed  in  10  per  cent  formalin  (1  part  of  commercial  formalin  to  9  parts  of  dis- 
tilled water)  and  left  indefinitely.  As  a  last  resort,  if  no  other  fixing  reagent 
is  available,  the  embryo  may  be  placed  in  the  strongest  alcohol  which  can 
be  secured  and  later  transferred  to  80  per  cent  alcohol  for  preservation. 

The  specimen  should  not  be  handled  nor  allowed  to  lie  in  water.  When 
the  proper  reagents  are  not  at  hand,  carefully  wrap  the  object  in  cloth  and 
keep  it  on  ice  if  possible  until  they  can  be  secured.  Very  small  embryos 
may  be  fixed  and  preserved  with  membranes  intact;  older  ones  (6  weeks  to 
3  months)  should  have  the  membranes  ruptured.  To  secure  the  best  fixa- 
tion of  fetuses  (2  months  and  beyond),  the  specimen  should  be  divided,  or  at 
least  the  body  cavity  should  be  opened. 

15.  For  Micro-Dissection  of  Small  Embryos,   after  fixation  Heuser 
stains  for  24  hours  in  alum-cochineal  diluted  with  5  times  its  volume  of  water. 
He  then  fixes  the  embryo  to  a  small  square  of  thin  ground  glass  with  celloidin 
cement  (about  0.75  per  cent  solution  of  celloidin)  and  dissects  in  alcohol 


Some  Embryological  Methods  135 

under  the  binocular  microscope.  The  dissected  specimen  may  be  preserved 
at  any  stage  by  placing  it,  still  attached  to  its  glass  support,  in  alcohol  in  a 
shell  vial  of  suitable  size.  Necessary  data  may  be  written  with  a  pencil 
on  the  ground  surface. 

Streeter  (American  Journal  of  Anatomy,  IV,  No.  1,  p.  87),  after  dehy- 
drating in  absolute  alcohol  attaches  the  embryo  with  a  drop  of  thick  celloidin 
to  a  smoked  isinglass  strip  coated  with  thin  celloidin  and  places  it  in  80  per 
cent  alcohol.  The  black  serves  as  a  good  background  for  the  embryo  and 
may  be  written  upon.  During  dissection  the  isinglass  strip  is  clamped  to  a 
glass  slide  which  has  been  cemented  with  balsam  to  one  facet  of  a  cut-glass 
polyhedral  paper-weight.  The  object  can  thus  be  placed  in  any  desired 
plane  and  dissected  in  alcohol  under  a  binocular  microscope. 

16.  Artificial  Fecundation  when  it  can  be  practiced  is  the  most  convenient 
means  of  securing  early  stages  of  development.  This  is  possible  with  many 
worms,  coelenterates,  echinoderms,  cyclostomes,  teleosts,  and  anuran 
amphibia. 

a)  In  echinoderms  (e.g.,  sea  urchin)  the  female  is  cut  open  and  a  number 
of  the  living  eggs  transferred  to  a  watch-glass  which  contains  fresh  sea  water. 
The  testes  of  a  male  are  teased  out  in  sea  water  and  a  drop  of  the  mixture 
is  conveyed  by  means  of  a  pipette  into  the  dish  containing  eggs.    Imme- 
diately upon  fertilization  a  membrane  forms  around  each  fertilized  egg.    In 
about  40  to  50  minutes  after  fertilization  the  signs  of  the  first  cleavage  should 
appear.    The  blastula  forms  in  about  6  hours,  and  the  gastrula  in  about  12 
hours.     For  the  study  of  fertilization,  etc.,  the  following  stages  should  be 
fixed  in  Bouin's  fluid  (p.  29)  for  30  minutes  and  stained  in  iron-hematoxylin 
(p.  51);   5  minutes  after  fertilization,  nucleus  giving  off  polar  bodies;   30 
minutes  after  fertilization,  approaching  pronuclei;   50  to  55  minutes  after 
fertilization,  division  of  nucleus  (mitotic  figure)  in  the  first  cleavage. 

b)  In  amphibia  (e.g.,  frog)  both  male  and  female  are  cut  open,  the  vasa 
deferentia  or  testes  are  teased  out  in  a  watch-glass  full  of  water,  and  the  ova 
are  then  removed  from  the  lower  ends  of  the  oviducts  and  placed  in  this  water. 
After  fertilization  the  eggs  should  be  placed  in  glass  dishes  in  not  over  4 
inches  of  water.    Many  eggs  should  not  be  placed  in  one  dish.    See  also 
memoranda,  pp.  121-123. 

c)  In  teleosts  the  eggs  are  obtained  by  stripping  the  female  when  she 
is  in  spawning  condition.    At  such  times  the  eggs  are  loose  in  the  body 
cavity  and  may  be  pressed  out  by  gently  manipulating  the  belly  of  the  fish. 
The  head  of  the  fish  should  be  held  in  one  hand,  the  tail  in  the  other,  and  the 
thumb  or  the  thumb  and  forefinger  used  to  press  out  the  ova  into  a  clean, 
dry  finger-bowl.    The  milt  of  the  male  is  obtained  in  the  same  manner  in 
a  dish  containing  a  little  fresh  or  sea  water,  depending  upon  the  habitat  of  the 
fish.    When  the  water  becomes  milky  with  sperm  pour  the  mixture  over  the 
eggs.    Eggs  and  sperm  are  then  gently  stirred  about  by  means  of  a  feather 


136  Animal  Micrology 

to  insure  thorough  mixing.  However,  in  some  teleosts  (e.g.,  stickleback, 
Fundulus)  it  is  necessary  to  kill  the  male  and  tease  out  the  testes.  In  the 
cunner  (Ctenolabrus)  10  minutes  after  fertilization  the  formation  of  blasto- 
disk  and  polar  bodies  may  be  observed;  30  to  33  minutes  after  fertilization 
the  two  pronuclei  may  be  found  in  close  approximation. 

If  other  than  the  very  early  stages  are  required,  the  fertilized  eggs  must 
be  transferred  to  a  hatching-box  or  jar,  depending  upon  the  kind  of  egg. 
This  is  best  done  by  means  of  a  horn  spoon  and  a  feather.  Dead  eggs, 
recognizable  by  their  opacity,  should  be  removed  at  least  once  a  day.  The 
conditions  under  which  the  eggs  of  different  species  thrive  are  so  varied  that 
the  reader  must  be  referred  for  details  to  such  special  publications  as  those 
of  the  United  States  Bureau  of  Fisheries  or  the  fish  commissions  of  the 
various  states. 

17.  For  the  Study  of  Early  Cleavage  in  Living  Material  the  eggs  of  some 
of  the  water  snails  afford  an  abundance  of  excellent  material.    By  watching 
aquaria  which  contain  snails  the  fresh  material  can  easily  be  obtained  dur- 
ing the  spring  and  summer.    Twigs  and  bits  of  board  to  which  the  egg-masses 
may  be  attached  should  be  placed  in  the  aquaria. 

If  one  is  at  the  seashore,  the  sea  urchin,  starfish,  squid,  various  marine 
annelids,  mollusks,  and  coelenterates  afford  an  abundance  of  material.  Of 
the  marine  fishes  Fundulus  and  Ctenolabrus  are  excellent.  Of  fresh-water 
fishes  the  whitefish  (spawning  in  November  or  December)  and  the  pickerel 
(in  April)  show  cleavage  well,  although  in  the  whitefish  it  is  very  slow. 

18.  For  Quick  Preparation  of  Cleavage  Stages  for  study  in  toto,  in  forms 
where  there  is  considerable  yolk,  Spaeth  finds  useful  often  a  mixture  of  equal 
parts  of  glacial  acetic  acid,  glycerin,  and  water,  to  which  enough  Delafield's 
hematoxylin  is  added  to  make  it  a  light  tan  color. 

19.  Chinese  Black  added  to  the  water  on  the  slide  in  which  eggs  with 
very  transparent  jelly  (e.g.,  Nereis)  are  being  examined  outlines  the  egg 
distinctly  and  shows  the  path  of  the  spermatozoon  through  the  jelly. 

20.  For  the  Study  of  the  Formation  of  Polar  Bodies,  Fertilization,  and 
Early  Cleavage  in  Sections  nothing  surpasses  the  eggs  of  Ascaris.    The 
Ascaris  (A.  megalocephala)  from  the  horse  is  preferable,  although  A.  lumbri- 
coides  from  the  pig  will  answer. 

The  ovisacs,  two  in  number,  are  very  long  convoluted  tubes.  Differ- 
ent regions  contain  eggs  in  different  stages  of  development.  The  thicker 
tubes  toward  the  anterior  end  of  the  animal  contain  cleavage  stages;  back 
of  these  are  cells  showing  extrusion  of  the  polar  bodies  and  fertilization 
stages.  The  material  must  be  fresh;  either  bring  the  live  Ascaris  to  the 
laboratory  or  take  the  fixing  fluid  to  the  place  for  obtaining  the  material. 
Slit  open  the  abdominal  wall  of  the  worm  and  remove  the  ovisacs  and  after 
separating  the  numerous  convolutions  somewhat,  fix  them  entire  for  24  hours 
in  a  mixture  of  absolute  alcohol  4  parts,  glacial  acetic  acid,  1  part,  or  for 


Some  Embryological  Methods  137 

15  to  25  minutes  in  acetic-alcohol-chloroform  (reagent  2,  6,  p.  207)  saturated 
with  corrosive  sublimate.  Preserve  in  80  per  cent  alcohol.  To  locate 
eggs  of  the  desired  stage  tease  out  eggs  at  intervals  along  the  ovisacs,  stain 
with  acid  carmine  (reagent  38,  p.  222),  and  examine.  The  proper  region  once 
located,  cut  out  small  lengths  of  the  tube,  imbed  :t  in  paraffin,  and  make 
thin  transverse  sections.  In  order  to  keep  the  eggs  from  shriveling,  the  bath 
in  hot  paraffin  must  be  curtailed.  Use  the  method  for  delicate  objects  (p.  53) . 
Stain  by  the  iron-hematoxylin  method  (p.  51).  Ascaris  eggs  when  smeared 
on  a  slide  in  thick  albumen  fixative,  which  is  then  coagulated  with  forma- 
lin, will  go  on  developing  if  put  into  an  incubator. 

21.  The  Cultivation  of  Removed  Embryonic  Tissues  in  clotted  lymph, 
plasma,  nutrient  agar,  bouillon,  and  in  various  salt  solutions  is  an  important 
phase  of  embryological  technique  which  has  been  developed  largely  in  the 
past  ten  years,  but  the  subject  is  too  extensive  to  treat  in  detail  in  limited 
space.  A  general  method  is  given  and  the  reader  is  left  to  look  up  modifica- 
tions and  other  methods  in  the  papers  listed  at  tl^e  end  of  this  memorandum. 

All  dissections  must  be  carried  on  under  aseptic  conditions.  Sterilize 
all  instruments,  pipettes,  slides,  covers,  and  vaseline  in  a  Bunsen  flame. 
For  cultivation  of  tissues  use  a  Locke's  solution  to  which  dextrin  has  been 
added.  It  is  made  as  follows: 

To  100  c.c  of  distilled  water  add: 

NaCl 0 . 900  per  cent 

CaCl2, 0.025  per  cent 

KC1 0.042  per  cent 

NaHCO3, 0.020  per  cent 

Dextrin 0.250  per  cent 

Remove  an  8-  or  10-day  chick  embryo,  under  aseptic  conditions,  to  about 
10  or  20  c.c.  of  the  sterilized  solution  heated  to  39°  C.  Cut  out  bits  of  intes- 
tine, kidney,  liver,  heart,  or  spleen  a  few  millimeters  in  diameter  and  place 
into  another  dish  which  contains  10  to  20  c.c.  of  the  solution  at  39°  C.  Cut 
each  small  piece  up  into  smaller  pieces  a  fraction  of  a  millimeter  thick. 
Draw  these  up  into  a  sterilized  fine  pipette,  one  at  a  time,  with  some  of  the 
solution  and  make  hanging-drop  preparations  (Fig.  40)  of  them  on  sterile 
cover-slips  which  are  thoroughly  clean  and  free  from  every  trace  of  grease. 
Invert  each  cover-slip  on  to  a  vaseline-ringed,  hollow-ground  slide  which  has 
been  sterilized.  For  rings  use  a  vaseline  melting  at  about  46°  C.  A  bit 
of  paraffin  may  be  added  to  ordinary  vaseline  to  stiffen  it. 

Incubate  the  cultures  at  about  39°  to  40°  C.  Growth  begins  within  10 
to  20  hours  and,  as  indicated  by  the  number  of  mitotic  figures,  reaches  its 
maximum  on  the  second  or  third  day.  When  it  is  desired  to  examine  the 
living  tissue  do  so  on  a  warm  stage.  The  margins  of  the  growing  regions 
are  the  best  points  to  examine  because  there  the  cells  are  only  one  or  two 
layers  thick.  (Method  of  Lewis  and  Lewis.) 


138  Animal  Micrology 

When  permanent  preparations  are  desired  the  cover-slip  is  removed  from 
the  vaseline  ring  and  the  film  of  tissue  is  fixed,  on  the  cover-slip,  by  means 
of  osmic-acid  vapor.  After  fixation  the  denser  central  piece  of  tissue  is 
torn  away,  leaving  only  the  thin  film  of  new  growth  which  is  treated  on  the 
cover-slip  as  one  treats  sections  on  a  slide.  Stain  in  iron-hematoxylin  or  in 
Ehrlich's  hematoxylin  and  eosin. 

Further  details  of  tissue  culture  in  vitro  and  bibliographies  will  be  found 
in  the  following  papers:  Harrison,  Anatomical  Record,  I  (1907);  Journal  of 
Experimental  Zoology,  IX  (1910);  Carrel  and  Burrows,  Journal  of  Experi- 
mental Medicine,  XIII  (1911);  Lewis  and  Lewis,  Johns  Hopkins  Hospital 
Bulletin,  XXII  (April,  1911),  241;  Anatomical  Record,  VI,  Nos.  1  and  5 
(1912);  American  Journal  of  Anatomy,  XVII,  No.  3  (March,  1915);  Anatomi- 
cal Record,  X,  No.  4  (February,  1916). 

22.  The  Living  Embryo  of  the  Chick  may  be  kept  under  observation 
for  some  hours  while  still  in  the  egg  by  employing  one  of  the  so-called  "win- 
dow" methods.  The  simplest  method  is  to  cut  out  a  disk  of  the  shell  on  one 
side  under  as  nearly  aseptic  conditions  as  possible,  so  that  the  embryo  is 
exposed.  A  bit  of  the  white  is  removed  and  a  film  of  celloidin  is  laid  over  the 
opening  to  form  the  window.  It  must  adhere  firmly  at  every  point  around 
the  margin.  The  embryo  will  continue  to  develop  for  some  time  if  the  egg 
is  put  back  into  the  incubator. 


CHAPTER  XVII 
SOME  CYTOLOGICAL  METHODS 

In  the  very  many  cytological  methods  which  have  been  in  vogue 
during  the  past  few  years  two  fixing  fluids,  Flemming's  strong  and 
Bourn's,  and  two  stains,  iron-hematoxylin  and  safranin,  stand  pre- 
eminent as  of  general  utility.  Iron-hematoxylin  with  or  without 
a  counterstain  may  be  used  successfully  after  either  of  the  fixing 
fluids  mentioned.  The  safranin  is  more  likely  to  prove  successful 
after  Flemming's,  although  in  some  materials  good  preparations  can 
be  made  with  it  after  Bouin's  fluid.  Two  other  fixing  fluids,  Gilson's 
mercuro-nitric  and  the  acetic-sublimate  mixture  of  Carnoy  and 
Lebrun,  are  also  of  wide  application,  especially  when  followed  by 
iron-hematoxylin  as  a  stain. 

A  recent  method  of  fixation  developed  by  Dr.  Ezra  Allen  gives 
promise  of  equaling  if  not  surpassing  any  of  the  foregoing,  particularly 
in  the  study  of  mammalian  tissues.  The  method  is  given  on  p.  149. 

In  preparing  tissues  for  cytological  work  it  is  imperative  that 
pieces  should  be  small,  not  more  than  3  or  4  mm.  thick  where  prac- 
ticable, to  insure  thorough  and  even  fixation.  To  secure  penetration 
from  all  sides,  it  is  well  to  place  a  few  layers  of  filter-paper  in  the 
bottom  of  the  vessels  in  which  tissues  are  fixed  and  to  shake  the  tissues 
about  a  little  from  time  to  time.  Mechanical  injury  may  be  avoided 
by  binding  a  bit  of  clean  linen  on  the  ends  of  the  forceps  with  which 
tissues  are  handled. 

I.    MITOSIS 

For  general  study  of  cell  structures,  and  particularly  cell  division, 
I  have  found  nothing  which  can  readily  be  obtained  in  quantities 
sufficient  for  class  use  that  surpasses  the  crayfish  testis,  the  blastodisk 
of  the  whitefish,  the  epidermis  and  testis  of  Ambystoma  and  Necturus, 
and  the  maturation  and  cleavage  stages  of  Ascaris.  Ascaris  material 
has  already  been  discussed  (p.  136). 

139 


140  Animal  Micrology 

Robertson  (Journal  of  Morphology,  XVII  [June,  1916])  finds  that 
in  grasshoppers  of  the  family  Tettigidae,  taken  before  the  last  moult, 
cells  dividing  mitotically  may  be  found  in  large  numbers  in  the  mesen- 
teron,  proctodaeum,  fat-body,  hypodermis,  and  the  follicles  of  the 
gonads.  The  columnar  epithelium  of  the  mesenteron  seems  to  be 
the  most  favorable  region  for  finding  such  divisions.  Inasmuch  as 
the  members  of  this  family  have  only  thirteen  or  fourteen  chromo- 
somes, the  material  should  prove  to  be  exceptionally  valuable  for 
purposes  of  class  demonstration 

Testis  of  Crayfish 

The  testicular  cells  of  the  crayfish  (Cambarus  virilis)  will  be 
found  in  active  proliferation  from  the  middle  of  June  to  the  middle 
of  July.  The  chromosomes  are  too  small  and  too  numerous  for 
satisfactory  individual  study,  but  the  spindles  and  centrosomes  are 
distinct  and  the  general  pictures  of  representative  stages  are  clear-cut 
and  easily  found. 

Section  Method. — 1.  Fix  small  bits  of  the  testes,  3  to  4  mm. 
thick,  in  Flemming's  fluid  (12,  p.  21 1)  for  24  hours.  Wash  in  running 
water  6  to  12  hours,  dehydrate,  and  imbed  in  paraffin  according  to  the 
methods  for  delicate  objects  (p.  53)  or  the  ''drop"  method  (6,  p.  152). 

2.  Cut  sections  5  to  7  microns  thick  and  mount  several  slides  by 
the  water-albumen  method. 

3.  After  removing  the  paraffin  with  xylol  and  running  the  slides 
down  through  the  alcohols,  stain  some  of  the  sections  by  the  iron- 
hematoxylin  method  (p.  51)  and  counterstain  with  acid  fuchsin  or 
orange  G.     Run  the  slides  up  through  the  alcohols,  clear  in  xylol, 
and  mount  in  a  thin  balsam.     Use  a  No.  1  cover-slip  if  the  prepara- 
tion is  to  be  studied  with  an  oil-immersion  lens. 

4.  Place  others  of  the  slides  in  safranin  (72,  p.  234)  for  24  hours, 
then  rinse  in  water  and  run  up  through  the  alcohols  to  95  per  cent. 
Counterstain  for  30  seconds  in  a  0 . 5  per  cent  solution  of  light  green 
(Lichtgrun  S.F.)  in  95  per  cent  alcohol.     If  the  sections  are  left  too 
long  in  the  green  stain  the  safranin  will  be  washed  out.     Pass  the 
slides  through  absolute  alcohol  into  clove  oil  for  a  few  minutes, 
rinse  in  xylol,  and  mount  in  thin  balsam  under  a  No.  1  cover-slip. 


Some  Cytological  Methods  141 

Smear  Preparations. — Remove  the  fresh  testis  to  a  slide  and 
tease  somewhat  with  needles  in  order  to  rupture  the  cysts  which 
inclose  the  germ  cells.  Spread  the  mass  evenly  over  the  slide  with 
the  end  of  another  slide  and  then  flatten  it  between  the  two  slides 
in  a  very  thin  film.  Avoid  any  considerable  pressure.  Separate 
the  slides  by  slipping  them  apart.  Each  should  bear  a  very  thin 
coating  of  the  material.  Plunge  them  into  Flemming's  fluid  and 
leave  for  24  hours.  Wash  in  running  water  for  6  to  12  hours,  then 
with  forceps  pick  or  scrape  off  all.  lumps  of  tissue  which  might  later 
keep  the  cover-slip  from  fitting  closely.  Stain  and  mount  as  if  the 
films  were  sections. 

If  preferred,  some  of  the  slides  can  be  fixed  in  Bourn's  fluid  for  an 
hour  or  two,  washed  in  50  per  cent  alcohol,  then  stained  in  iron- 
hematoxylin  and  counterstained  in  acid  fuchsin  or  orange  G. 

Blastodisk  of  Whitefish  (Coregonus) 

1.  Spawn  the  females  and  fertilize  the  eggs  (in  early  December) 
as  directed  in  memorandum  16,  p.  135. 

2.  Select  eggs  in  the  32-  to  64-cell  stage  of  cleavage  (40  to  60 
hours  after  fertilization)  and  fix  for  6  or  8  hours  in  Bouin's  fluid. 

3.  Wash  in  repeated   changes  of  50  per  cent  alcohol,  then  in 
70  per  cent  alcohol  until  the  yellow  color  ceases  to  come  from  the 
eggs. 

4.  With  needles  carefully  dissect  off  the  blastodisks  under  a 
binocular  or  other  dissecting  microscope. 

5.  Dehydrate,  section  in  paraffin  (method,  p.  36),  stain  in  iron- 
hematoxylin  with  or  without  a  counterstain,  and  mount  as  usual. 
Sections  should  be  about  7  microns  thick. 

The  eggs  of  the  pickerel,  obtainable  in  April,  may  be  handled 
with  equal  ease.  They  cleave  much  more  rapidly  than  do  those  of 

the  whitefish. 

Testis  of  Necturus 

The  cells  of  the  testes  will  ordinarily  be  found  undergoing  rapid 
proliferations  in  late  July  and  early  August.  Those  toward  the 
posterior  end  of  the  testis  show  the  most  advanced  stages  of  spermato- 
genesis,  those  toward  the  anterior  end  the  least  advanced  stages. 


142  Animal  Micrology 

Both  cells  and  chromosomes  are  very  large.  The  spindle  usually 
shows  up  well  and  the  chromosomes  exhibit  considerable  variety  in 
shape  and  size. 

1.  From  different  regions  of  the  testis  fix  some  bits  of  testis  in 
Bouin's  fluid  (6  to  8  hours)  and  others  in  Flemming's  (24  to  36  hours). 
Wash  out  the  Bouin  as  in  3,  p.  141,  and  the  Flemming  according  to 
directions  in  step  1,  p.  140. 

2.  Dehydrate,  imbed,  and  section  according  to  the  usual  paraffin 
method. 

Iron-Hematoxylin  Preparations. — 3.  Stain  sections  of  each  kind 
of  material  according  to  the  ordinary  iron-hematoxylin  method  with 
or  without  a  counterstain  (pp.  51-52.). 

Safranin-Gentian-Violet  Preparations. — 4.  Also  stain  some  of  the 
Flemming  material  according  to  the  safranin  and  gentian-violet 
method  (73,  p.  234). 

Safranin-Gentian-Orange  Preparations. — -Use  saturated  aqueous 
solutions  of  safranin,  gentian  violet,  and  orange  G  respectively. 
Rinse  and  stain  in  gentian  violet  2  to  5  minutes  (time  determined 
by  trial).  Pipette  absolute  alcohol  over  the  sections  until  the 
violet  is  out  of  the  cytoplasm,  then  follow  with  orange  G,  pipetting  it 
on  and  removing  it  again  almost  instantly.  Wash  off  with  absolute 
alcohol,  dip  in  oil  of  cloves,  clear  in  xylol,  and  mount  in  thin  balsam 
under  a  No.  1  cover. 

Somatic  Cells  of  Ambystoma 

•  • 

Epidermal  Cells. — Cut  off  the  tails  of  several  one-month-old 
Ambystoma  larvae  into  Flemming's  fluid.  At  the  end  of  2  to  4 
hours  strip  off  bits  of  the  epidermis  from  the  tails  and  fix  these 
strips  for  some  20  hours  longer  in  the  fluid.  Wash  in  running 
water  6  to  8  hours,  stain  some  according  to  the  iron-hematoxylin 
method  (p.  51)  and  others  with  safranin  and  light  green  (step  4, 
p.  140).  Dehydrate  and  mount  as  usual. 

If  Bouin's  is  used  instead  of  Flemming's  fluid,  the  peeling  off  of  the 
epidermis  need  not  be  done  until  the  end  of  fixation  (6  to  8  hours). 

Peritoneal  Cells. — Parmenter  recommends  larger  larvae  than 
those  used  for  epidermis.  He  cuts  away  the  side  walls  of  the  body 


Some  Cytological  Methods  143 

cavity,  pulls  out  the  intestine,  and  fixes  the  remaining  tissue  in  the 
region  of  the  spinal  column  in  Flemming's  or  in  Bouin's  fluid  as 
above.  Bits  of  the  peritoneum  on  either  side  of  the  dorsal  mid-line 
are  stripped  off  and  prepared  as  was  the  epidermis. 

Either  of  these  kinds  of  preparations  shows  splendid  polar  views 
of  cell-division  stages,  though  little  or  nothing  of  lateral  views. 
They  are  especially  favorable  for  showing  longitudinal  splitting  of 
chromosomes. 

Living  Cells. — Curarize  Ambystoma  or  other  young  amphibian 
larvae  by  adding  (according  to  size  of  larva)  5  to  10  drops  of  a  0.5 
per  cent  solution  of  curare  in  equal  parts  of  glycerin  and  water  to  a 
watch-glassful  of  water.  After  40  minutes  remove  for  half  an  hour 
to  a  1  per  cent  solution  of  sodium  chloride  in  water.  Wrap  in  blotting 
paper  and  examine  the  tail  fin  on  a  slide  under  the  microscope.  Cell 
divisions  may  be  seen  in  progress. 

If  replaced  in  fresh  water  such  larvae  recover  after  some  hours. 
After  curarization  some  workers  prefer  to  cut  the  larva  in  two  parts 
in  front  of  the  hind  limbs,  studying  only  the  tail.  The  gills  also 
show  interesting  cell  activities.  If  curare  is  not  at  hand  a  3  per 
cent  alcohol  or  ether  may  be  used,  although  not  so  successfully. 

H.    MITOCHONDRIA 

Recent  studies  tend  to  show  that  mitochondria  occur  more  or  less  exten- 
sively in  nearly  all  kinds  of  tissue.  They  were  largely  overlooked  in  the  past 
because  many  of  the  fixing  fluids  in  use  contain  strong  organic  acids,  such  as 
acetic,  and  these  dissolve  mitochondria.  They  are  sometimes  stained  with 
great  sharpness  by  iron-hematoxylin,  following  fixation  in  Flemming's 
strong  mixture  (12,  p.  211)  in  which,  instead  of  1  c.c.,  only  3  to  6  drops  of 
glacial  acetic  acid  are  used  for  every  15  c.c.  of  chromic  acid.  For  their 
careful  study,  however,  cytologists  are  using  special  methods.  These  are 
too  numerous  and  complex  to  be  reviewed  in  an  elementary  guide  to  tech- 
nique. Bibliographies  and  discussion  of  the  technical  details  will  be  found 
in  the  publications  of  Bensley  and  particularly  of  Cowdry.  See  Bensley, 
American  Journal  of  Anatomy,  XII  (1911),  297-388;  Cowdry,  Internationale 
Monatsschrift  fur  Anatomie  und  Physiologie,  XXIX  (1912);  American 
Journal  of  Anatomy,  XVII,  No.  1  (November,  1914);  ibid.,  XIX,  No.  3 
(May,  1916);  Contributions  to  Embryology,  No.  11,  Carnegie  Institution  of 
Washington. 


144  Animal  Micrology 

Three  methods  of  wide  application  are  as  follows: 
Benda's  Method. — 1.  Fix  for  eight  days  in  a  modified  Flemming  fluid 
made  as  follows: 

Chromic  acid,  1  per  cent 15  c.c. 

Osmic  acid,  2  per  cent 4  c.c. 

Glacial  acetic  acid 3  drops 

2.  Wash  in  water  for  1  hour,  then  for  24  hours  in  a  mixture  of  100  parts 
pyroligneous  acid  and  1  part  chromic  acid. 

3.  Transfer  to  a  2  per  cent  potassium  bichromate  solution  for  24  hours, 
run  up  through  the  grades  of  alcohol  to  xylol,  and  finally  infiltrate  with 
paraffin  and  section.    Sections  should  be  about  5  microns  thick. 

4.  After  removal  of  paraffin  from  sections  run  them  down  to  distilled 
water  and  place  them  in  a  4  per  cent  iron-alum  solution  for  24  hours. 

5.  Wash  thoroughly  in  water  and  transfer  to  a  solution  of  Kahlbaum's 
sulphalizarinate  of  soda  (made  by  taking  1  part  of  a  saturated  aqueous  solu- 
tion of  the  stain  to  from  80  to  100  parts  of  distilled  water)  for  24  hours. 

6.  Rinse  the  slide  in  distilled  water  and  flood  it  with  a  crystal  violet 
anilin-water  solution  (equal  parts  of  anilin  water  and  a  3  per  cent  solution  of 
the  dye  in  95  per  cent  alcohol).    Warm  until  the  solution  steams,  keeping 
it  heated  for  about  3  minutes. 

7.  Wash  in  distilled  water,  transfer  to  30  per  cent  acetic  acid  for  1  or  2 
minutes,  then  wash  in  running  water  for  5  or  10  minutes. 

8.  Dry  the  slide  with  filter  paper,  dip  it  for  a  minute  into  absolute 
alcohol,  place  in  oil  of  bergamot  until  cleared,  then  transfer  it  through  xylol 
and  mount  in  balsam  in  the  usual  way. 

A  successful  preparation  should  show  chromatic  elements  a  deep  purple 
and  the  cytoplasm  a  light  red  with  mitochondria  violet. 

Wildman  (Journal  of  Morphology,  XXIV,  No.  3  [1913])  modifies  the 
method  by  transferring  slides  from  the  alizarin  solution,  after  rinsing,  into 
a  3  per  cent  solution  of  crystal  violet  (3  c.c.  of  anilin  stain  in  100  c.c.  of  dis- 
tilled water)  for  10  minutes;  rinsing  and  passing  into  80  per  cent  alcohol 
for  5  seconds;  passing  through  95  per  cent  and  absolute  alcohol;  and,  when 
properly  differentiated,  clearing  and  mounting  in  the  usual  way. 

Bensley's  Acid-Fuchsin,  Methyl-Green  Methods. — Fix  tissues  for  24 
hours  in  the  following: 

Osmic  acid,  2  per  cent 2  c.c. 

Potassium  bichromate,  2 . 5  per  cent 8  c.c. 

Glacial  acetic  acid 1  drop 

Sections  should  be  4  microns  or  less  in  thickness.  Mount  by  the  water 
method  (p.  23),  remove  paraffin  with  toluol,  then  pass  through  absolute 
alcohol  to  water.  Treat  for  from  30  seconds  to  1  minute  (determined  by 


Some  Cytological  Methods  145 

trial)  with  a  1  per  cent  solution  of  potassium  permanganate,  then  for  the 
same  length  of  time  with  a  5  per  cent  solution  of  oxalic  acid.  The  per- 
manganate extracts  the  mordanting  elements  of  fixation  and  the  oxalic 
acid  removes  the  permanganate.  Thoroughly  wash  in  water. 

Stain  for  5  minutes  in  Altmann's  acid  fuchsin  (acid  fuchsin  20  grams, 
anilin  water  100  c.c.)  which  has  previously  been  warmed  to  60°  C.  Wash 
thoroughly  in  distilled  water,  dip  for  an  instant  into  a  1  per  cent  solution  of 
methyl  green,  then  wash,  rapidly  dehydrate  in  absolute  alcohol  (avoiding 
alcohols  of  intermediate  strengths),  clear  in  toluol,  and  mount  in  balsam. 
Toluidin  blue  can  be  substituted  for  methyl  green. 

If  the  material  does  not  stain  well  with  the  acid  fuchsin,  or  if  the  methyl 
green  or  toluidin  blue  obliterates  it,  treat  the  sections  with  a  2.5  per  cent 
aqueous  solution  of  potassium  bichromate  for  about  half  a  minute  and  rinse 
in  water  just  before  staining  in  acid  fuchsin. 

In  spinal  ganglion  cells,  for  example,  mitochondria  should  appear  bright 
red;  Nissl  substance,  green  (or  blue);  neurofibrils  in  the  axon  hillock,  light 
brown;  and  the  canalicular  systems  should  be  revealed. 

Bensley's  Copper-Chrome-Hematoxylin  Method. — Fix  materials  hi 
acetic-osmic-bichromate  mixture  and  prepare  for  staining  as  in  the  preceding 
method.  Wash  for  1  hour  in  distilled  water,  then  thoroughly  dehydrate. 
Leave  in  absolute  alcohol  for  24  hours,  then  pass  through  equal  parts  of 
bergamot  oil  and  absolute  alcohol  (1  hour)  into  pure  bergamot  oil  for  3 
hours,  followed  by  equal  parts  of  bergamot  oil  and  paraffin  (1  hour),  then 
by  paraffin  melting  at  60°  C.  (2  to  3  hours).  Cut  sections  4  microns  thick 
and  fix  to  the  slide  by  the  water  method.  Remove  paraffin  with  toluol  and 
pass  down  through  the  alcohols  to  distilled  water. 

Place  sections  for  5  minutes  in  a  saturated  aqueous  solution  of  copper 
acetate,  wash  in  several  changes  of  water,  and  transfer  for  a  minute  to  a 
0 . 5  per  cent  aqueous  solution  of  a  well-ripened  hematoxylin.  Wash  hi  water 
and  transfer  for  1  minute  to  a  5  per  cent  aqueous  solution  of  neutral  potassium 
chromate.  The  sections  should  turn  a  blue-black  color.  If  they  are  of  only 
a  light-blue  shade,  place  them  again  in  the  copper  acetate  and  repeat  the 
operations  from  there  on. 

Wash  several  minutes  in  water,  then  differentiate  under  the  microscope 
hi  Weigert's  borax-ferricyanide  mixture  (borax,  2  parts;  ferricyanide  of 
potassium,  2.5  parts;  water,  200  parts)  diluted  with  2  volumes  of  water. 
Wash  6  to  8  hours  in  tap  water,  then  dehydrate,  clear  hi  toluol,  and  mount 
in  balsam.  Mitochondria  should  appear  a  bluish  black  against  a  clear  back- 
ground. 

Mitochondria  in  Living  Cells  stain  specifically  with  Janus  green.  There 
are  several  Janus  greens,  but  of  these  Cowdry  reports  that  only  Janus  green 
B  (diethylsafraninazodimethylanilin  chloride)  of  the  Farbwerke  Hoechst 
Co.  (obtainable  from  L.  A.  Metz  &  Co.,  New  York)  will  give  the  desired 


146  Animal  Micrology 

reaction.  He  finds  that  mitochondria  will  stain  in  human  lymphocytes  in  a 
dilution  of  Janus  green  in  normal  saline  solution  of  1 : 500,000.  Ordinarily, 
for  living  tissue,  a  dilution  of  1 : 15,000  or  1 : 20,000  in  normal  saline,  in  Locke's 
solution,  or  in  Ringer's  solution,  is  employed.  Janus  blue,  G  and  R,  may  also 
be  used  as  a  vital  stain  for  mitochondria. 

While  fresh  tissues  may  be  stained  by  immersion  in  the  dye,  much  better 
results  are  obtained  by  injection  through  the  blood  vessels,  in  normal  saline, 
after  the  vessels  have  been  thoroughly  flushed  out  with  normal  saline  solu- 
tion. 

Mitochondria  in  Tissue  Grown  "in  Vitro"  (see  p.  137)  may  be  studied 
readily,  according  to  M.  R.  and  W.  H.  Lewis  (American  Journal  of  Anatomy, 
XVII,  No.  3  [1915]),  who  observed  their  changes,  growth,  and  division  in 
embryonic  tissues  of  the  chick. 

m.    STAINING  OF  LIVING  OR  FRESH  TISSUES 

Intra-vitam  staining,  so  called,  has  come  more  and  more  into  prominence 
during  the  past  few  years.  It  is  questionable  if  a  staining  of  really  "vital" 
elements  ever  occurs,  although  undoubtedly  various  granules  in  cells  may 
be  stained  while  the  cells  are  yet  living.  Certain  stains  also  may  be  used  with 
fair  success  with  fresh  or  lightly  fixed  cells. 

For  intra-vitam  staining,  neutral  red,  Bismarck  brown,  Janus  green,  and 
methylen  blue  are  the  dyes  most  commonly  employed.  They  are  used  in 
the  proportion  of  about  1  part  of  the  dye  to  10,000  or  20,000  parts  of 
some  normal  fluid,  such  as  normal  saline,  Ringer's  solution,  or  Locke's 
solution. 

For  lightly  fixing  and  staining  fresh  cells  methyl  green  acidulated  to 
about  0 . 75  per  cent  with  acetic  acid  is  in  common  use.  Also  for  the  study  of 
fresh  cells  tissues  are  teased  in  a  solution  of  Ripart  and  Petit  (p.  211)  to 
which  0. 1  per  cent  osmic  acid  has  been  added,  and  then  stained  in  methyl 
green.  Acid  carmine  is  frequently  used  for  the  study  of  chromosomes  in 
fresh  cells  (see  memorandum  2,  p.  151).  For  the  use  of  Sudan  III  with 
living  animals  see  p.  147.  Trypan  blue  will  make  certain  parts  of  the  living 
body  take  on  an  intense  blue  color,  but  the  color  seems  to  be  due  wholly 
to  the  engulfment  of  the  colored  particles  by  certain  phagocytic  cells, 
particularly  in  the  connective  tissues  of  the  body  (Evans  and  Schule- 
mann,  Science,  XXXIX  [1914],  443-54).  Thus,  1  c.c.  of  a  0.5  per 
cent  solution  injected  into  the  peritoneal  cavity  of  a  mouse  will  rapidly 
blue  it  from  ears  to  tail  without  noticeably  interfering  with  its  normal 
activities. 

Equal  parts  of  glycerin,  95  per  cent  alcohol,  and  distilled  water  is  a 
useful  examining-medium  in  which  fresh  tissues  may  be  kept  for  a  long  time 
without  deterioration. 


Some  Cytological  Methods  147 

IV.    TESTS  FOR  CERTAIN  CELLULAR  STRUCTURES 

The  following  tests  while  not  always  specific  are  serviceable  in  helping 
to  identify  some  of  the  more  usual  cellular  contents: 

Archoplasm  stains  intensely  with  acid  fuchsin  or  light  green. 

Calcification  may  usually  be  detected  by  means  of  3  to  5  per  cent  hydro- 
chloric acid.  When  treated  with  this  solution  carbonate  of  lime  emits 
bubbles  of  carbon  dioxide,  while  phosphate  of  lime  simply  dissolves. 

Cell  Walls  are  usually  well  denned  by  acid  fuchsin  when  used  as  a  counter- 
stain  with  some  of  the  hematoxylins. 

Centrosomes  are  best  shown  by  the  iron-hematoxylin  long  method 
(p.  52).  Heidenhain  finds  that  they  are  more  sharply  defined  if  the  sec- 
tions, previous  to  mordanting  in  iron-alum,  are  stained  for  24  hours  in  a  weak 
solution  of  Bordeaux  red. 

Chromatin. — In  fresh  cells,  methyl  green  stains  only  chromatin  when  it 
colors  any  part  of  the  cell.  Absence  of  coloration  with  this  dye  does  not 
necessarily  mean  absence  of  chromatin.  Moderate  digestion  with  gastric 
jaice  (at  about  40°  C.)  will  remove  albumins  and  leave  chromatin.  Pro- 
longed treatment  with  1  per  cent  caustic  potash  or  with  fuming  hydrochloric 
acid  will  remove  all  chromatin  from  a  nucleus.  A  10  per  cent  solution  of 
sodium  chloride  swells  chromatin  and  may  dissolve  it. 

After  sublimate  fixations,  thin  (3-micron)  sections  stained  by  the  Ehrlich- 
Biondi  method  (41,  p.  222)  or  Auerbach's  fuchsin-methyl-green  method 
(46,  p.  224)  should  show  "active"  chromatin  or  chromosomes  green,  linin, 
and  plasmosomes  red. 

Chromosomes. — The  best  single  stain  in  fixed  material  is  either  iron- 
hematoxylin  or  safranin,  although  chromatoid  bodies  and  mitochondria, 
when  present,  may  also  stain  by  these  reagents.  In  fresh  tissue,  acid  carmine 
(38,  p.  222)  colors  chromosomes,  as  does  also  methyl  green  (60,  p.  231). 

Fat. — As  the  fat  of  tissues  is  dissolved  by  xylol,  alcohol,  and  other  rea- 
gents used  in  the  paraffin  and  celloidin  methods,  only  teased,  free-hand,  or 
frozen-sectioned,  fresh  material,  or  material  fixed  in  some  non-fat  solvent 
fixer  such  as  formalin  or  Mtiller's  fluid,  can  be  used. 

Osmic  Acid  (21,  p.  215)  is  the  commonest  test  for  fat.  It  stains  most  but 
not  all  fatty  bodies  brown  or  black.  Osmicated  fats  are  rendered  sufficiently 
insoluble  to  permit  of  dehydration  and  mounting  in  balsam,  if  absolute  alcohol 
is  avoided  and  cedar  oil  is  used  instead  of  xylol  for  clearing.  However,  euparal 
(p.  54)  may  be  used. 

Sudan  III  is  a  specific  stain  for  fat  (see  76,  p.  235).  Large  fat  drops  stain 
from  a  brilliant  red  to  an  orange;  small  ones  may  be  yellowish  red.  The  fat  of 
animals  fed  with  the  dye  will  become  intensely  colored  by  it.  The  fat  in  the 
layers  of  yolk  laid  down  in  hens'  eggs  while  the  fowls  are  fed  on  the  dye  (Riddle, 
Science,  XXVII  [1908],  945)  is  stained  red,  and  eggs  so  colored  (Gage)  hatca  into 
chicks  with  the  body  fat  colored  pink. 


148  Animal  Micrology  " 

Scharlach  R  is  superseding  Sudan  III  as  a  stain  for  fat.  It  stains  fat  orange 
to  red.  If  a  permanent  mount  is  desired,  frozen  sections  may  be  fixed  10  minutes 
in  formalin  vapor,  stained  for  12  hours  in  a  saturated  filtered  solution  of  the  dye 
in  70  per  cent  alcohol,  washed  in  water,  counterstained  in  alum-hematoxylin, 
washed  and  mounted  in  glycerin  or  glycerin-jelly.  However,  see  74,  p.  235. 

Fat  may  be  removed  from  tissues  ordinarily  by  treatment  with  alcohol,  ether, 
or  chloroform. 

Free  Acid  in  tissues  may  be  detected  by  Congo  red,  the  solutions  of 
which  become  blue  in  presence  of  free  acid.  Neutral  red  is  turned  bright 
red  by  acid,  yellow  by  alkalies. 

Lecithin  may  be  distinguished  from  fat  by  its  less  solubility  in  ether  and 
its  greater  capacity  for  stains.  Formalin-fixed  material  if  brought  into 
acetone  has  the  fat  dissolved,  but  not  its  lecithin.  The  latter  may  be  stained 
by  osmic  acid,  hematoxylin,  orange  G,  acid  fuchsin,  methyl  green,  or  toluidin 
blue,  although  the  tissue  should  be  dehydrated  in  acetone  and  left  as  little 
as  possible  in  alcohol. 

Glycogen. — Readily  soluble  in  aqueous  media,  hence  tissues  should  be 
fixed  and  hardened  in  95  per  cent  alcohol.  Gage  (The  Microscope,  p.  278) 
states  that  a  Lugol's  solution  made  of  1 . 5  grams  of  iodine  crystals,  3  grams 
of  iodide  of  potassium,  1.5  grams  of  sodium  chloride,  and  300  c.c.  of  water 
gives  a  differential  stain  (a  mahogany  red)  for  glycogen  in  sections.  For 
very  soluble  glycogen  he  recommends  that  50  per  cent  alcohol  be  substituted 
for  the  water  in  the  stain.  He  deparaffins  with  xylol,  mounts  in  yellow  vase- 
line, and  seals  with  shellac  or  balsam. 

Hemoglobin,  after  proper  fixation,  stains  a  characteristic,  clear,  deep-red 
color  with  eosin.  For  crystals  see  p.  105. 

Intra-cellular  Reduction  Processes  may  be  detected  by  Janus  green  used 
as  an  intra-vitam  stain  (p.  146).  With  reduction  the  color  changes  from 
blue  or  green  to  red. 

Mitochondria  (see  pp.  143-146). — The  most  nearly  specific  single  stain 
for  mitochondria  in  fresh  tissue  is  probably  Janus  green  (p.  145). 

Mucin  in  cells,  after  sublimate  fixation,  stains  with  basic  but  not  with 
acid  anilin  dyes.  Either  thionin  or  toluidin  blue  stains  mucin  reddish, 
surrounding  elements  blue.  Methylen  blue  and  safranin  are  also  good  stains 
for  mucin.  See  also  muci-carmine  and  muci-hematin  (62  and  63,  p.  231). 

Nissl's  Granules  (tigroid  substance). — For  the  methylen-blue  method 
see  p.  230.  See  also  pp.  145  and  232. 

Oxidase  Reaction  (Schultze's). — The  presence  of  an  oxidizing  ferment 
in  cells  may  be  disclosed  by  the  following  method : 

Solution  1. — Boil  I  gram  of  a-naphthol  in  100  c.c.  of  distilled  water  until  it 
melts.  Add  pure  potassium  hydrate  (about  1  c.c,.)  until  the  naphthol  is  com- 
pletely dissolved.  The  solution  should  pass  from  yellow  to  yellowish  brown. 


Some  Cytological  Methods  149 

Solution  2. — Make  a  1  per  cent  aqueous  solution  of  dimethyl-p-phenylendiamin 
(Merck)  at  room  temperature.  Filter. 

Use  frozen  sections  of  formalin-fixed  material  or  cover-glass  preparations  fixed 
in  vapor  of  formaldehyde.  Move  the  preparations  gently  back  and  forth  in  so- 
lution 1  for  about  3  minutes,  then  do  the  same  in  solution  2.  Wash  in  distilled 
water  and  mount  and  examine  in  water  or  in  glycerin-jelly.  Oxidase  granules 
are  stained  deep  blue. 

Plasmosomes  remain  unstained  in  fresh  material  treated  with  acid 
methyl  green  (60,  p.  231)  which  stains  chromatin.  With  the  Ehrlich- 
Biondi  stain  they  stain  red,  sometimes  orange;  with  safranin,  gentian  violet, 
and  other  basic  dyes,  in  regressive  staining  (p.  24),  the  plasmosomes  retain 
the  stain  more  tenaciously  than  resting  chromatin  does.  Because  of  greater 
refractivity  they  are  frequently  demonstrable  in  unstained  preparations  and 
sometimes  in  living  cells. 

Secretion  Antecedents. — See  Bensley,  American  Journal  of  Anatomy, 
XII,  No.  3  (1911);  XIX,  No.  1  (1916). 

Spindle  Fibers  are  frequently  well  stained  by  acid  fuchsin  when  used  as 
a  counterstain  after  such  fixers  as  Flemming's,  Gilson's,  or  Bouin's  fluids. 
See  also  "Euparal,"  p.  54. 

V.    ALLEN'S  METHOD  (Anatomical  Record,  X,  No.  9  [July,  1916]) 
Fix  tissues  in  Allen's  B-15  fluid,  which  is  made  up  as  follows: 

Picric  acid,  saturated  aqueous  solution 75  c.c. 

Formalin  (c.p.) 25  c.c. 

Glacial  acetic  acid 5  c.c. 

Just  before  using  heat  to  37°  C.  and  add  1 . 5  grams  of  chromic  acid 
crystals,  agitating  the  mixture  vigorously  until  the  crystals  are  dissolved. 
Then  add  2  grams  of  urea  crystals.  During  fixation  keep  the  liquid  heated 
to  37°  or  38°  C. 

Pieces  of  brain  0.5  c.c.  in  volume  fix  in  1  hour.  Bits  of  young  mature 
testes  require  a  little  longer  and  pieces  of  older  testes  2  to  3  hours. 

Let  the  fluid  and  the  contained  tissue  cool  to  room  temperature  and 
dehydrate  gradually  by  the  drop  method  (memorandum  6,  p.  152)  with 
alcohol  up  to  75  per  cent  alcohol,  then  finish  dehydration  with  anilin  oil. 
Regulate  to  about  one' drop  per  second,  or  less  if  the  quantity  of  fixing  fluid 
is  small.  Bring  pieces  of  soft  tissue,  some  0.5  c.c.  in  volume,  up  to  75  per 
cent  alcohol  in  about  1  hour.  Harder  tissues  require  more  time. 

To  wash  out  all  picric  acid  thoroughly,  replace  the  ordinary  75  per  cent 
alcohol  on  the  object  with  75  per  cent  alcohol  containing  a  few  drops  of  a 
saturated  aqueous  solution  of  lithium  carbonate.  Keep  up  agitation  with  a 
very  slow  curre'nt  of  air  and  continue  the  washing  until  the  yellow  color 


150  Animal  Micrology 

ceases  to  appear  in  the  fluid.  As  soon  as  possible  after  washing,  to  avoid 
shrinkage  in  alcohol,  start  replacement  by  anilin  oil,  letting  it  drop  in  slowly. 
To  insure  rapid  mixing  a  stronger  current  of  air  may  be  required;  when 
nearly  pure  anilin  is  reached  leave  the  tissue  in  it  until  it  is  clear  like  amber. 

Replace  the  anilin  with  bergamot  oil  or  synthetic  oil  of  wintergreen, 
following  the  same  method.  Change  the  oil  once  after  the  tissue  has  arrived 
in  pure  oil. 

Warm  the  oil  and  tissue  slightly  and  add  to  it  every  10  minutes  a  few 
drops  of  melted  paraffin,  which  must  be  thoroughly  mixed  with  the  oil  by 
means  of  a  pipette.  When  the  mixture  is  about  85  or  90  per  cent  paraffin, 
transfer  the  object  to  pure  paraffin  with  a  melting-point  of  52°  to  55°  C. 
If  bergamot  oil  has  been  used,  make  at  least  four  changes  of  paraffin.  Leave 
the  tissue  in  each  about  30  minutes  and  in  a  fifth  paraffin  about  an  hour. 
Testis  material  requires  longer  time.  Imbed  and  section  in  the  usual  way. 

VI.    PHOTOGRAPHING  CELLULAR  STRUCTURES 

• 

1.  Select  a  slide  in  which  the  part  to  be  photographed  is  well  stained  in 
iron-hematoxylin.    The  background  should  be  unstained,  as  a  sharp  con- 
trast in  the  slide  gives  better  results  in  the  picture. 

2.  Using  an  apochromatic  lens  and  a  compensating  or  projection  ocular, 
bring  the  part  selected  for  the  picture  into  sharp  focus  under  the  microscope. 

3.  Place  the  microscope  containing  the  object  in  focus  under  the  camera. 
Adjust  the  bellows  to  any  desired  length.     Remember  that  a  lower  ocular 
with  a  longer  bellows,  at  the  same  magnification,  produces  a  better  picture 
than  a  higher  ocular  with  a  shorter  bellows.    Also  remember  that  in  using 
a  camera  mounted  in  a  vertical  plane  there  is  less  likelihood  of  jarring 
the  object  out  of  focus  when  the  microscope  is  placed  under  the  camera  than 
when  it  is  adjusted  in  a  horizontal  plane. 

4.  Focus  the  object  under  the  microscope  upon  the  ground-glass  screen 
of  the  camera.    To  insure  sharp  focus,  a  focusing  glass  should  be  used.    The 
picture  may  now  be  taken. 

5.  Foot   and   Strobell    (Zeitschrift  fur   wissenschaftliche  Mikroskopie, 
XVIII  [1901],  421-26)  developed  the  following  method,  which  obviates 
focusing  upon  the  ground  glass  of  the  camera  every  time  a  picture  is  taken. 
Procure  from  an  oculist  several  concave  spectacle  lenses  (—1  to  —10  diop- 
ters).   When  steps  1  to  4  have  been  completed,  remove  the  microscope 
from  under  the  camera  without  the  least  shift  of  focus.    Do  not  touch  the 
micrometer  or  fine  adjustment  screws  of  the  microscope.     Place  over  the  ocular 
one  after  another  of  the  spectacle  lenses  until  one  is  found  through  which  the 
object  appears  in  as  sharp  focus  as  it  did  upon  the  ground-glass  screen. 
Note  the  number  of  the  particular  lens  used;    Thereafter,  when  an  object 
is  to  be  photographed  with  the  same  ocular  and  objective,  the  same  tube  and 


Some  Cytological  Methods  151 

bellows  length,  one  need  only  place  the  dioptric  lens  over  the  ocular, 
focus  until  the  points  desired  in  the  finished  photograph  stand  out  sharply, 
remove  the  dioptric  lens,  place  the  microscope  under  the  camera  without* 
shift  of  focus,  and  take  the  picture.  Since  the  dioptric  lens  corrects  the  focus 
for  that  bellows  length,  the  necessity  of  refocusing  upon  the  ground  glass  is 
removed.  Whenever  a  different  combination  of  ocular  objective  and  bellows 
length  is  used,  the  proper  dioptric  lens  must  be  found. 

6.  It  is  good  practice  to  allow  the  microscope  to  stand  for  a  time  after 
the  object  has  been  focused  through  the  dioptric  lens  in  order  to  be  sure 
that  the  focus  does  not  shift.  After  the  picture  has  been  taken,  by  replacing 
the  lens  over  the  ocular  one  can  see  if  the  focus  has  been  held  throughout 
the  time.  Even  with  the  utmost  care  the  focus  will  sometimes  change. 

MEMORANDA 

1.  Accessory  Chromosomes  (sex  chromosomes,  ^-elements)  are  perhaps 
best  demonstrated  in  the  testes  of  some  species  of  the  short-horned  grass- 
hoppers taken  about  the  time  of  the  last  moult.    In  these  forms  mitotic 
figures  are  large  and  chromosomes  usually  distinct.    Among  the  Hemiptera, 
the  squash  bug  (Anasa  tristis)  for  single  X-elenient,  and  the  stink  bug 
Euschistus,  for  X-  and    F-elements,  are  recommended.    Flemming's  or 
Bouin's  fluid  may  be  used  for  fixing,  and  iron-hematoxylin  or  safranin  for 
staining. 

2.  For  Quick  Determination  of  the  Chromosomal  Condition  of  cells 
aceto-carmine  preparations  (38,  p.  222)  are  useful.    The  entire  testes  of  an 
insect  is  put  on  the  slide  in  a  drop  of  the  stain  and  the  cells  separated  by  slight 
pressure  on  the  cover,  or  smears  from  the  testes  of  larger  animals  are  made  and 
flooded  with  the  stain,  after  which  a  cover-slip  is  added.    The  edges  of  the 
cover  should  be  sealed  with  vaseline  to  prevent  evaporation.    Chromosomes 
are  stained  in  a  few  minutes.    Such  preparation  should  be  carefully  checked 
by  observations  on  well-fixed  and  stained  materials,  however,  since  the  great 
amount  of  acetic  acid  in  the  acid  carmine  swells  chromosomes  and  is  likely 
to  lead  to  erroneous  conclusions  regarding  details. 

3.  Protoplasmic  Currents  in  cells  may  be  seen  to  good  advantage  in 
Rhizopods,  in  the  plasmodia  of  Myxomycetes,  and  in  the  stamens  of  Trades- 
cantia. 

4.  Celloidin  Instead  of  Paraffin  is  being  used  more  and  more  by  various 
cytologists  for  making  sections,  the  purpose  being  to  avoid  the  bad  effects 
of  hot  paraffin.    For  example,  Danchakoff  (Zeitschrift  fur  wissenschaftliche 
Mikroskopie,  XXV  [1908])  starts  with  very  thin  celloidin,  changes  to  some- 
what thicker,  then  to  still  thicker  (about  3  per  cent),  and  finally,  having 
arranged  the  tissues,  lets  the  solvent  evaporate  very  slowly  throughout  4 
or  5  days  to  a  week.    The  mass  should  become  opaline,  homogeneous,  and 


152 


Animal  Micrology 


about  as  hard  as  vulcanized  rubber.    It  is  stored  in  80  per  cent  alcohol. 
Thin  sections  can  be  cut  if  the  mass  is  sufficiently  hard. 

5.  Urea  is  being  used  very  successfully  in  such  fixing  fluids  as  Flemming's 
and  Bouin's  by  Professor  C.  E.  McClung  and  his  pupils  (see  Allen's  method, 
p.  149).    From  1  to  3  or  more  grams  per  100  c.c.  of  fixer  is  the  quantity  used. 
The  exact  proportions  for  any  particular  tissue  can  be  determined  only  by 
trial. 

6.  Very  Gradual  Changing  of  Fluids  is  recommended  in  the  treatment 
of  tissues  to  be  used  for  cytological  studies.    This  may  be  accomplished  very 


ji. 

H  Waste  H 


Drying  bottle 


FIG.  42.  —  Apparatus  for  Gradual  Change  of  Liquids  (after  Ezra  Allen) 

successfully  according  to  the  drop  method  described  by  Allen  in  the  Anatomi- 
cal Record  for  July,  1916.  The  apparatus  he  designed  for  the  purpose  is 
shown  in  Fig.  42. 

A  2-  or  3-liter  aspirator  bottle  (W.B.)  is  filled  with  water  and  its  stop- 
cock opened  slightly  until  the  water  begins  to  drop  into  the  tightly  corked 
pressure  bottle  (P.B.).  As  the  air  is  compressed  in  this  bottle  bubbles  will 
begin  to  issue  from  the  air  tube  through  the  liquid  in  the  small  container 
((7)  in  which  the  tissue  lies.  To  secure  a  steady  stream  of  air  through  C, 
the  end  of  the  air  tube  is  drawn  out  into  almost  a  capillary  and  the  rubber 
tube  connecting  it  with  P.  B.  is  clamped  nearly  shut.  The  purpose  of  the 
current  of  air  is  to  insure  quick  and  thorough  mixing  of  liquids  in  C. 

The  alcohol  or  other  replacing  fluid  in  the  supply  bottle  (S.B.)  is  started 
to  dropping  at  the  desired  rate,  the  flow  being  regulated  by  a  stop-cock  or 


Some  Cytological  Methods  153 

clamp.    A  siphon  on  the  side  of  C  removes  the  excess  of  fluid  into  a  waste 
jar,  so  that  the  concentration  of  the  liquid  which  is  being  added  steadily 


For  higher  alcohols  or  oils  the  air  should  be  dried  by  passing  it  through 
a  tube  con  taming  calcium  chloride  or  through  sulphuric  acid  (see  figure). 
Two  liters  of  water  in  W.B.  should  last  all  night  at  the  rate  of  a  drop  a  second. 

7.  The  Use  of  Anilin  Oil  in  Place  of  the  Higher  Alcohols  for  completing 
dehydration  is  strongly  advocated  by  Allen  (op.  tit.,  p.  149)  because  delicate 
tissues  are  less  likely  to  shrink  in  it.    Since  anilin  oil  does  not  mix  with 
paraffin  it  must  be  followed  by  some  clearing  oil.    Allen  prefers  oil  of  berga- 
mot  or  synthetic  oil  of  wintergreen. 

8.  Cooling  Tissues  by  placing  them  on  ice  for  15  or  20  minutes  before 
fixation  may  sometimes  prove  helpful  in  preventing  the  clumping  or  sticking 
together  of  chromosomes.    With  difficult  material,  such  as  that  of  birds 
or  mammals,  it  is  well,  at  least,  to  try  this  method  along  with  others. 

9.  The  Mature  Testes  of  Young  Mammals  or  Birds  are  better  for  studies 
in  spermatogenesis  than  those  of  older  animals. 

10.  For  Dissection  of  Living  Cells  an  apparatus  designed  by  Barber 
is  being  used  very  successfully  by  several  investigators.    It  is  described  in 
full  in  the  Kansas  University  Science  Bulletin,  IV  (March,  1907)  and  in  the 
Philippine  Journal  of  Science,  IX  (1914).     Some  workers  employ  a  double 
form  of  it. 

11.  For  the  Estimation  of  Very  Minute  Quantities  of  Carbon  Dioxide, 
Tashiro  has  designed  several  pieces  of  accurate  and  delicate  apparatus 
which  are  described  in  full  in  the  American  Journal  of  Physiology,  XXXII, 
No.  2  (1913),  107-45,  and  the  Journal  of  Biological  Chemistry,  XVI,  No.  4 
(1914),  485-94.    These  various  types  of  apparatus  may  be  obtained  from 
the  Eimer  &  Amend  Co.,  of  New  York  City. 

12.  Holmgren's  Canals  may  be  demonstrated  in  various  cells  by  fixing 
in    Bensley's   formal-bichromate-sublimate   mixture     (Biological  Bulletin, 
XIX,  No.  3  [August,  1910]).    The  composition  of  the  fluid  is  as  follows: 

Neutral  formalin  (freshly  distilled) 10  c.c. 

Water 90  c.c. 

Potassium  bichromate 2.5  grams 

Mercuric  chloride 5      grams 

Use  the  solution  soon  after  making  up. 

13.  Euparal  (VIII,  p.  54)  is  of  great  value  in  the  study  of  certain  achro- 
matic cellular  elements. 


CHAPTER  XVIII 
RECONSTRUCTION  OF  OBJECTS  FROM  SECTIONS 

In  investigating  objects  which  possess  complex  internal  cavities 
or  complicated  structures  it  is  frequently  very  difficult  to  gain  an 
adequate  idea  from  the  direct  study  of  serial  sections,  or  by  means  of 
macerated  or  teased  preparations;  consequently  various  methods 
of  plastic  or  geometrical  reconstruction  from  the  sections  are  resorted 
to.  For  such  reconstruction,  sections  must  be  of  uniform  thickness, 
serial,  and  they  must  possess  similar  orientation. 

RECONSTRUCTION  IN  WAX 

Bern's  method  of  constructing  wax  models  of  objects  from  serial 
sections  is  widely  used  for  both  embryological  and  anatomical  sub- 
jects. The  thickness  of  the  sections,  the  magnification  of  the  micro- 
scope, and  the  plane  of  section  must  be  known. 

Wax  plates  are  prepared  as  many  times  thicker  than  the  actual 
sections  as  the  latter  will  be  magnified  in  diameters.  For  example, 
if  the  serial  sections  are  -gV  of  a  millimeter  thick  (33 J  microns),  and 
they  are  to  be  magnified  60  diameters,  then  the  wax  plates  must  be 
made  60  times  as  thick  as  the  sections,  or  2  mm.  thick.  This  is  a 
thickness  commonly  used.  Count  the  number  of  sections  to  be 
reconstructed,  and  prepare  an  equal  number  of  plates. 

Preparation  of  the  Wax  Plates 

a)  The  hot-water  method. — 1.  Prepare  the  wax  according  to  the  following 
formula: 

Beeswax 6  parts 

Paraffin  (melting-point  56°  C.) 4  parts 

White  lump  (not  powdered)  rosin 2  parts 

Melt  together  and  thoroughly  mix. 

2.  To  prepare  plates  of  the  proper  thickness  (2mm.),  use  shallow 
straight-walled  rectangular  tin  pans  which  wiU  afford  a  water  surface  of 
3X4  feet;  2,000  grams  of  the  prepared  wax  poured  on  very  hot  water  in 

154 


Reconstruction  of  Objects  from  Sections  155 

such  a  pan  will  give  a  plate  2  mm.  thick.  Air  bubbles  which  form  in  the 
wax  may  be  driven  off  before  it  cools  by  playing  a  Bunsen  flame  over  the 
surface.  The  wax  should  spread  evenly  over  the  surface  of  the  water  if 
both  wax  and  water  are  sufficiently  hot.  If  gaps  remain,  close  them  by 
drawing  a  glass  slide  over  the  surface  of  the  wax.  To  prevent  the  plate  from 
splitting  while  cooling,  after  it  has  stiffened  somewhat,  cut  the  edges  free 
from  the  walls  of  the  pan.  When  the  water  has  become  tepid,  remove  the 
wax  plate  to  a  flat  support  and  leave  it  to  harden. 

6)  The  wax-plate  machine  method. — Several  instruments  have  been 
devised  for  making  the  plates  more  rapidly  and  more  accurately  than  by  the 
hot-water  method.  Huber's  apparatus,  for  instance,  consists  of  a  heavy 
cast-iron  plate  with  movable  side  pieces  which  can  be  adjusted  to  a  height 
corresponding  to  the  desired  thickness  of  the  wax  plates.  The  whole  instru- 
ment is  supported  upon  three  adjustable  legs,  by  means  of  which  it  can  be 
made  exactly  level.  Melted  wax  slightly  in  excess  of  the  quantity  necessary 
for  a  wax  plate  is  poured  on  to  the  iron  plate  in  an  even  layer,  and  rolled 
out  with  a  hot  roller  until  the  roller  comes  to  run  directly  on  the  side  pieces 
of  the  instrument.  When  the  wax  plate  is  cool  enough  to  handle,  it  may  be 
placed  in  a  pan  of  cold  water  to  harden. 

Practical  Exercise. — When  possible  an  outline  drawing  of  the 
part  to  be  reconstructed  should  be  made  before  it  is  sectioned. 

1.  Reconstruct  the  heart  of  a  chick  at  the  end  of  the  third  day 
of  incubation,  under  a  magnification  of  60  diameters.     For  this 
magnification,  if  it  is  desired  to  use  a  wax  plate  2  mm.  thick,  the 
original  sections  should  have  been  33 . 3  microns  thick. 

2.  Place  a  sheet  of  blue  tracing-paper  on  the  wax  plate  with  the 
colored  side  toward  it.     Over  the  tracing-paper  place  a  sheet  of 
ordinary  drawing-paper.     With  the  aid  of  a  camera  lucida  or  other 
projection  apparatus,  outline  on  the  drawing-paper  the  part  to  be 
reconstructed.     In  doing  this  the  outline  is  also  traced  in  blue  on  the 
wax.    Number  each  drawing,  and  also  indicate  the  number  of  the 
section  on  the  slide  to  which  it  corresponds;   also  number  the  wax 
plates  with  reference  to  the  drawings. 

3.  Lay  the  wax  plate  on  a  suitable  flat  surface,  and  cut  out  the 
outlined  parts  with  a  sharp,  narrow-bladed  knife.     Leave  bridges 
of  wax  to  hold  in  place  the  parts  that  would  otherwise  be  separate 
pieces.     Pile  up  the  successive  sections  in  proper  sequence  as  they 
are  cut  out. 


156  Animal  Micrology 

4.  In  finally  putting  the  model  together,  accurately  adjust  the 
parts  (for  reconstruction  points  see  memorandum  13,  II,  a,  p.  127), 
and  build  up  the  model  in  blocks  of  five  sections  each  (Bardeen's 
suggestion).  If  necessary,  unite  the  essential  parts  by  means  of 
pins  or  fine  nails.  Remove  all  temporary  wax  bridges  (see  3)  by 
means  of  a  hot  knife. 

When  all  blocks  are  properly  adjusted  and  united,  smooth  over 
the  surface  by  means  of  a  hot  spatula. 

MEMORANDA 

1.  Geometrical  Reconstructions,  first  described  by  Professor  His,  are 
often  all  that  is  necessary  to  give  one  the  desired  information  about  internal 
organs.  Before  sectioning,  an  outline  drawing  of  the  object  is  made  in  a 
plane  at  right  angles  to  the  intended  plane  of  section,  and  under  the  same 
magnification  that  will  be  used  for  the  reconstructed  drawing.  For  example, 
if  the  sections  are  to  be  transverse,  the  outline  drawing  of  the  object  would 
be  a  profile  view  from  the  side.  After  sectioning  the  object,  each  section 
is  drawn  under  the  same  magnification  as  was  used  for  the  outline  drawing. 

To  reconstruct  any  special  part  of  the  object,  draw  a  median  line  on 
the  outline  drawing  corresponding  to  the  long  axis  of  the  object.  At  right 
angles  to  this  line  draw  a  series  of  equidistant  parallel  lines  corresponding 
in  positions  to  the  sections  that  have  been  made.  For  example,  if  the  magni- 
fication is  100  diameters  and  the  sections  10  microns  thick,  then  the  parallel 
lines  must  be  1  mm.  apart.  Then,  beginning  with  the  first  section,  indicate 
by  dots  in  the  proper  plane  in  the  profile  drawing  the  relative  distances  of 
the  part  in  the  sections  above  or  below  the  median  line  along  the  proper  one 
of  the  parallel  lines.  All  of  the  sections  having  thus  been  plotted,  connect 
the  dots  of  corresponding  parts  in  the  successive  zones.  It  is  frequently 
sufficient  to  reconstruct  only  every  fifth  or  even  every  tenth  section.  When 
the  plane  of  section  is  not  quite  at  right  angles  to  the  axis  of  the  object,  an 
equal  alteration  of  angle  must  be  made  between  the  median  line  of  the 
outline  drawing  and  the  parallel  lines. 

Such  a  reconstruction  as  that  above  would  give  lateral  views  of  the  various 
internal  parts.  To  get  their  aspects  as  seen  from  above  or  below,  the  original 
outline  drawing  of  the  specimen  as  a  whole  should  have  been  made  from 
this  point  of  view  instead  of  from  the  side.  In  actual  work  one  should  make 
reconstructions  in  both  planes. 

For  a  modification  of  Weber's  method  of  graphic  reconstruction,  see 
Scammon,  Anatomical  Record,  IX,  No.  3  (March,  1915).  For  suggestions 
on  profile  reconstructions  see  Streeter,  American  Journal  of  Anatomy,  IV, 
No.  1  (1904),  86. 


Reconstruction  of  Objects  from  Sections  157 

2.  A  Special  Drawing-Table  for  rapid  and  convenient  drawing  of  sections 
for  reconstruction  has  been  devised  by  Bardeen.    For  details,  see  Johns 
Hopkins  Bulletin,  XII,  148. 

3.  Sheets  of  Blotting  Paper  instead  of  wax  are  recommended  by  Mrs. 
Gage  (Anatomical  Record,  I,  No.  7  [November,  1907]).    Models  are  finished 
by  coating  with  paraffin.    The  advantages  claimed  for  this  method  are 
lightness,  durability,  and  ease  and  cleanliness  of  production.    The  method 
is  also  given  in  Gage,  The  Microscope,  pp.  326-32. 

4.  Plates  with  the  Paper  of  the  Drawing  Rolled  into  the  Wax,  following 
directions  in  Karl  Peter,  Methoden  der  Rekonstruktion  (Gustav  Fischer,  Jena), 
have  been  found  very  satisfactory  by  Rice.    A  thin  drawing-paper,  smooth 
on  the  drawing  side,  porous  on  the  other,  is  used  and  drawings  are  duplicated 
by  means  of  a  carbon  sheet.    One  copy  is  kept  for  reference,  the  other  is 
pressed  into  the  plate.    To  accomplish  the  latter,  the  slab  on  which  the  wax 
is  rolled  out  is  smeared  thoroughly  with  turpentine  and  the  drawing  laid 
face  down  on  it.    The  melted  wax  is  then  poured  upon  the  back  of  the  draw- 
ing and,  when  beginning  to  harden,  is  covered  with  tissue  paper,  which  is 
then  rolled  into  it.    The  two  papers  thus  present  good  surfaces  for  building 
up  the  plates.    The  thicker  paper  with  the  drawing  on  it  gives  a  fixed  contour 
which  is  a  helpful  guide  when  it  comes  to  smoothing  down  the  model. 

Twisted  wires  in  short  lengths  are  used  for  supports.  The  ends  are 
spread  to  afford  anchorage  where  they  lie  between  the  plates.  Two  longer 
wires  should  be  twisted  together,  then  cut  into  proper  lengths. 

5.  Photography  of  Sections  upon  Large  Plates  has  been  resorted  to  by 
Warren  H.  Lewis  (Anatomical  Record,  IX,  No.  9  [September,  1915],  719-29) 
as  a  substitute  for  the  laborious  and  time-consuming  method  of  drawing 
each  section  on  paper.    He  uses  line  bromide  or  azo  G  hard  (matte)  prints. 
He  considers  the  photographs  far  superior  to  drawings  and  maintains  that, 
when  time  is  taken  into  account,  the  method  is  less  expensive  than  the  old 
method  of  tracing.    Lewis'  paper  is  full  of  valuable  suggestions  and  should 
be  read  by  everyone  who  contemplates  doing  much  work  in  reconstruction. 
The  chief  points  emphasized  are:  the  use  of  photographs;  the  use  of  series 
of  guide-lines  which  coincide  with  planes  that  are  at  right  angles  to  each  other 
and  perpendicular  to  the  plane  of  the  sections;   and  the  use  of  plaster-of- 
Paris  casts. 

6.  For  Rapidly  Cutting  Out  the  Wax  Plates,  Chester  H.  Heuser,  of  the 
Wistar  Institute  of  Anatomy  and  Biology,  has  devised  a  series  of  metal 
styli  of  varied  design  which  when  electrically  heated  are  handled  much  as 
one  would  handle  a  pen  in  writing.    The  cutting  instrument  proper  (copper, 
iron,  or  brass  wire,  pointed  or  flattened  at  one  end  according  to  need)  is 
coated,  except  for  about  15  mm.  at  the  working  end,  with  a  thin  layer  of 
an  insulating  asbestos  paste.    A  piece  of  No.  32  German  silver  wire  about 


158  Animal  Micrology 

20  cm.  long  is  then  wrapped  around  the  coated  surface,  with  the  coils  kept 
well  insulated  from  each  other,  and  finally  covered  with  the  paste.  The 
ends  of  the  German  silver  wire  are  attached  to  small  copper  wires  which 
run  to  a  lamp-board  bearing  several  incandescent  bulbs  with  sockets  con- 
nected in  parallel.  The  lamp-board  serves  as  a  rheostat,  so  that  any  desired 
temperature  can  be  obtained  in  the  stylus  by  altering  the  number  of  lights 
and  thus  regulating  the  current  which  passes  through  the  German  silver 
wire.  Styli  of  different  kinds  of  metal  attached  in  common  to  the  same 
lamp-board  afford  different  temperatures  for  cutting  points  and  smoothers. 
Heuser  contemplates  inserting  small  rheostats  in  the  system  to  regulate  the 
temperatures  of  the  individual  instruments  more  accurately. 

Professor  Mark  (Proceedings  of  the  American  Academy  of  Arts  and 
Sciences,  XLII,  No.  23  [1907])  uses  an  electrically  heated  wire  moved 
rapidly  by  a  modified  sewing  machine  for  cutting  out  the  wax  plate. 


CHAPTER  XIX 

DRAWING 

I  should  make  it  absolutely  necessary  for  everybody,  for  a  longer  or 
shorter  period,  to  learn  to  draw.  It  gives  you  the  means  of  training  the  young 
in  attention  and  accuracy,  which  are  the  two  things  in  which  all  mankind 
are  more  deficient  than  in  any  other  mental  quality  whatever. — Huxley. 

Drawing  is  an  important  part  of  the  work  in  most  biological 
sciences.  The  essential  phases  of  a  subject  can  be  condensed  into  a 
few  pages  if  the  drawings  accurately  represent  the  dissections  or 
microscopical  preparations  studied.  The  following  simple  direc- 
tions are  written  mainly  to  aid  the  student  in  preparing  his  notebook, 
but  it  is  hoped  that  they  may  also  be  useful  to  individuals  preparing 
manuscripts  for  publication. 

Materials  for  Class  Work. — All  the  materials  needed  for  ordinary 
class  work  can  be  selected  from  the  list  here  given  with  the  approxi- 
mate price  attached. 

Pencils — one  4H,  one  HB,  one  2B,  10  cents  each. 
Pens — Gillott's  lithographic  penpoint,  No.  290,  5  cents  each. 
Ink — water-proof  India  ink  made  by  Charles  Higgins  &  Co.,  retails 
at  25  cents  per  bottle. 

Ruler — celluloid,  10  cents. 

Ruby  eraser — 10  cents. 

Crayon  pencils — red,  blue,  and  yellow,  5  cents  each. 

Loose-leaf  notebook  with  bond  paper,  40  cents. 

The  pencils  may  be  of  any  standard  make;  the  4H  is  a  hard 
pencil  for  line  work;  the  HB,  a  medium  pencil,  is  useful  in  placing 
outlines  and  shading.  The  2B,  for  black  shading,  should  not  be  used 
unless  the  drawings  are  afterward  fixed  to  prevent  rubbing.  The 
ruler  graded  in  centimeters  on  one  edge  and  inches  on  the  other  is 
indispensable.  The  eraser  is  for  erasing  pencil  lines;  for  erasing  ink 
a  sharp  knife  is  best.  A  larger  assortment  of  colored  pencils  will 
often  prove  useful,  but  the  three  primary  colors  will  answer  most 
purposes.  Two-ply  bristol  board  at  2  cents  per  sheet  may  be  used 

159 


160  Animal  Micrology 

in  the  notebook  instead  of  the  bond  paper.  Where  classes  are  large, 
bookstores  will  make  up  bound  notebooks  for  35  cents  each,  contain- 
ing a  good  grade  of  paper  upon  which  drawings  can  be  made.  Where 
pencil  drawings  only  are  required,  pens  and  ink  may  be  omitted  from 
this  list.  On  account  of  the  small  amount  of  locker  or  drawer  space 
usually  available  for  one  student,  an  elaborate  drawing  outfit  should 
be  avoided.  The  excellence  of  student  drawings  is  judged  by  the 
exactness  with  which  the  preparations  are  depicted. 

I.    METHODS  OF  REPRESENTATION 

Outline. — In  beginning  a  drawing,  the  field  which  the  picture  will 
occupy  should  be  marked  off  with  dots.  The  placing  of  two  faint 
lines  which  cross  at  right  angles  in  the  middle  of  the  drawing-field 
is  a  great  help  to  beginners,  especially  in  drawing  bilaterally  sym- 
metrical objects.  Next  determine  the  relation  of  the  length  of  the 
object  to  the  breadth;  then  calculate  the  size  of  the  drawing.  If  the 
object  is  large,  a  reduction  will  be  necessary;  if  small,  it  can  be 
represented  better  5  or  10  times  its  original  size.  The  important 
points  can  be  indicated  in  the  drawing-field  by  dots  which,  when  con- 
nected by  light  lines,  roughly  block  in  the  object  in  correct  proportion 
and  size.  This  crude  picture  may  then  be  worked  over  until  angles 
are  removed  and  a  neat  outline  results.  For  the  preliminary  mapping 
of  the  object  an  HB  pencil  is  best,  as  the  lines  are  easily  erased.  The 
outline  when  finished  must  consist  of  a  continuous  line  of  uniform 
thickness  with  no  overlapping  edges  where  the  pencil  has  been 
removed  from  the  paper  and  put  down  again.  Outline  is  the  most 
important  part  of  the  drawing,  for  "a  good  outline  may  redeem  bad 
finish,  but  no  amount  of  excellence  in  finish  can  save  a  picture  that 
has  been  incorrectly  outlined."  What  details  to  include  depends 
upon  the  purposes  of  the  drawing.  The  principal  points  of  an 
anatomical  drawing  stand  out  more  clearly  when  they  are  not 
obscured  by  unnecessary  details.  In  histological  drawings,  details 
are  essential,  but  they  should  still  be  kept  subordinate  to  the  general 
effect  of  the  picture. 

Depth. — Usually  the  third  dimension,  depth,  is  not  considered 
in  drawings  made  from  sections.  When  drawings  of  reconstructions 


Drawing  161 

or  whole  mounts  are  made,  however,  all  three  dimensions  must  be 
indicated  in  the  drawing.  Likewise,  drawings  of  such  things  as 
digestive  canal,  nerve  cord,  heart,  lungs,  and  kidneys  stand  out 
better  when  depth  is  represented.  This  can  usually  be  done  by 
indicating  degrees  of  light  and  shade. 

Ink  Drawings. — In  drawings  which  are  to  be  inked,  the  outline 
should  be  carefully  drawn  in  pencil  and  as  many  corrections  as 
possible  made  before  ink  is  applied.  Place  the  ink  upon  the  pen  by 
means  of  the  quill  attached  to  the  cork  of  the  ink-bottle.  If  the 
original  outline  is  even,  the  inking  can  be  done  readily;  a  line  uniform 
in  thickness  results  from  the  applications  of  firm,  steady  pressure. 
The  pen  will  give  a  ragged  line  if  held  so  that  one  nib  bears  more 
heavily  upon  the  paper  than  does  the  other,  or  if  it  becomes  sticky 
with  dried  ink.  A  smoother  line  will  be  obtained  if  the  penholder 
is  held  at  a  wide  angle  to  the  paper  and  only  the  very  point  of  the 
pen  is  allowed  to  touch. 

Shading. — Where  differentiation  of  parts  is  desired,  shading  may 
be  used.  This  can  be  done  either  by  stippling  or  by  lines.  In 
making  the  dots  in  a  stippled  drawing,  the  pen  must  be  grasped 
firmly  and  only  the  point  placed  upon  the  paper.  If  the  pen  strikes 
the  paper  at  an  acute  angle,  three-sided  instead  of  round  dots  result. 
The  dots  must  all  be  of  the  same  size.  To  indicate  degrees  of  shade, 
vary  the  number  of  dots,  not  their  size.  A  heavy  shading  can  be 
accomplished  by  placing  the  dots  close  together,  whereas  dots  farther 
apart  give  the  impression  of  light  shading.  Lines  can  be  used  with 
good  effect  upon  large  drawings.  Let  the  lines,  placed  an  even 
distance  apart,  follow  the  shape  of  the  shadows.  To  indicate  heavy 
shadows,  lines  in  an  opposite  direction  can  be  placed  across  the  first 
set.  Be  careful  not  to  cross-hatch  until  the  first  lines  have  dried, 
otherwise  blots  will  occur. 

Pencil  Drawings. — When  correctly  executed,  pencil  drawings 
are  more  artistic  and  permit  of  more  subtle  differentiation  in  detail. 
Minute  points  can  be  shown  by  the  use  of  stippling.  In  stippling  with 
a  pencil,  follow  the  same  procedure  as  in  the  use  of  a  pen.  The  pencil- 
point  should  be  sharp  and  rounded  on  all  sides.  In  large  drawings, 
lines  can  be  evenly  placed  to  outline  shadows,  but  they  are  not  as 


162  Animal  Micrology 

effective  as  shadows  put  in  by  blending  graphite,  obtained  by  rubbing 
the  pencil  over  the  paper.  For  the  latter  method  a  stub  is  necessary. 
This  can  be  made  from  a  strip  of  paper  1  inch  wide  and  5  inches  long 
in  the  following  manner:  Begin  to  roll  the  paper  at  one  end  and  let 
each  turn  overlap  the  preceding  turn  slightly,  until  an  elongated  coil 
results.  The  pointed  end  of  this  can  be  used  in  spreading  graphite 
evenly  over  a  surface.  The  graphite  is  placed  on  the  part  of  the 
drawing  where  the  darkest  shadows  occur  with  an  HB  or  2B  pencil, 
and  is  worked  over  with  the  end  of  the  stub  until  the  sharp  edges  of  the 
shadow  gradually  grade  out  into  the  lighter  parts.  The  2B  pencil 
ordinarily  should  not  be  used  as  the  source  of  the  graphite,  as  shadows 
can  be  darkened  by  repeating  the  application  of  the  HB  pencil. 

Shadows. — In  most  pictures  of  biological  subjects,  one  cannot 
stand  off  and  observe  where  the  light  falls  upon  the  object  and  what 
part  is  in  shadow.  For  that  reason  a  knowledge  of  where  the  shadows 
occur  if  an  object  is  illuminated  from  any  one  direction  is  necessary. 
To  gain  such  knowledge  from  a  description  is  impossible;  it  is 
therefore  advisable  for  students  wishing  to  shade  their  drawings 
to  consult  an  artist  who  can  give  usable  information  in  the  form  of 
demonstrations.  A  careful  study  of  textbook  illustrations  will  aid 
materially.  Shading  requires  practice,  and  even  then  it  may  not 
be  successful.  In  most  cases  where  not  imperative  it  had  better  be 
left  out  entirely. 

Fixing  pencil  drawings. — Where  pencil  drawings  are  made  with 
soft  pencils  which  are  liable  to  rub,  they  must  be  fixed.  This  is  done 
with  a  fixing  solution  and  a  special  atomizer  which  can  be  bought  at 
any  art  store.  To  prepare  the  fixative,  make  a  saturated  solution 
of  white  shellac  in  alcohol.  Allow  this  to  stand  for  a  day  or  so ;  dilute 
one-half;  then  filter  off  the  liquid.  To  prevent  evaporation  when  not 
in  use,  this  must  be  kept  in  a  tightly  stoppered  bottle. 

The  drawing  should  be  placed  in  an  upright  position,  about  2  feet 
from  the  spray.  In  order  to  avoid  a  glossy  surface  spray  lightly. 

Wash-Drawings.— After  a  faint  outline  of  the  section  or  object 
has  been  made  with  a  hard  pencil,  fasten  the  paper  to  a  board  with 
thumb  tacks,  and  with  a  large  brush  dampen  the  entire  surface, 
removing  the  surplus  water  with  the  brush  or  a  blotter.  Mix  the 


Drawing  163 

wash  as  follows:  With  a  wet  brush  remove  some  pigment  from  a 
cake  of  Winsor  &  Newton's  Charcoal  Grey  or  Ivory  Black,  and  put 
in  the  water  in  the  mixing-pan.  Repeat  this  process  until  the  wash 
is  slightly  darker  than  the  desired  background  tint.  Next  put  a  wash 
of  this  over  the  entire  background;  it  will  dry  lighter.  Allow  the 
paper  to  dry  partly  before  darker  tones  are  put  in.  Where  a  very 
dark  portion  is  confined  to  a  small  area,  the  paper  should  be  quite  dry, 
otherwise  the  wash  will  run  into  the  surrounding  part  of  the  drawing. 
If  details  are  to  be  put  in  by  stippling  or  linework,  make  a  wash  (from 
the  same  cake)  the  color  and  consistency  of  ink.  This  can  be  applied 
with  a  pen  or  brush  after  the  paper  dries. 

Some  artists  use  a  dry  paper  which,  however,  requires  more  skill 
in  applying  the  wash.  The  darkest  tones  can  be  applied  first,  gradu- 
ally working  up  to  the  lightest.  After  experimenting,  use  that  which 
gives  the  best  results  for  the  purpose  in  hand. 

Where  several  wash-drawings  are  to  be  made  with  the  same  tones 
it  will  be  found  simpler  to  put  in  all  the  backgrounds  first.  Mix  up 
plenty  of  wash  for  this  purpose,  as  it  is  not  easy  to  duplicate  the  exact 
shade  at  another  mixing.  Wash  allowed  to  stand  becomes  darker 
upon  evaporation  of  the  water;  hence,  if  after  the  backgrounds  are 
put  in  the  work  must  be  deferred  until  later,  the  same  wash  will  do 
for  darker  tints.  Do  not  redampen  the  whole  surface,  as  that  will 
lighten  the  background.  The  darker  tones  can  be  blended  into  the 
background  with  a  clean  wet  brush. 

MEMORANDA 

1.  Cleanliness. — Even  where  the  drawings  are  correctly  made  as  to  size 
and  proportion,  the  notebook  will  not  present  a  good  appearance  if  the  pages 
contain  finger-marks  and  blots  to  mar  their  whiteness.    With  sufficient 
diligence  finger-marks  can  be  erased,  but  the  best  way  is  not  to  make  them 
in  the  beginning.    Where  laboratory  work  requires  dissection,  rough  sketches 
can  be  made  upon  scrap  paper  and  later  copied  into  the  notebook.    Blots 
can  be  avoided  if  care  is  used  in  placing  the  ink  upon  the  pen;   a  quill  is 
attached  to  the  stopper  of  the  ink-bottle  for  this  purpose. 

2.  Size  and  Arrangement  of  Drawings. — Uniformity  in  size  and  arrange- 
ment of  drawings  should  be  preserved  wherever  possible.    For  example, 
in  the  development  of  the  frog's  egg,  no  increase  in  the  size  of  the  egg  takes 


164  Animal  Micrology 

place  from  the  single-cell  stage  to  the  end  of  gastrulation,  hence  all  drawings 
representing  this  series  of  development  must  be  the  same  size.  The  neural 
tube,  largest  in  circumference  in  the  head,  decreases  gradually  toward  the 
posterior  end  of  the  body,  and  must  be  drawn  correctly  in  cross-sections, 
otherwise  one  will  have  an  erroneous  idea  of  its  structure.  The  drawings 
must  not  be  crowded  upon  the  page.  Exact  margins  and  spaces  equi- 
distant between  drawings,  •  though  not  artistic,  give  the  impression  of  neat- 
ness desirable  in  scientific  work. 

Labeling. — The  results  will  not  equal  expectation  if  the  labeling  is 
poorly  done.  Too  few  students  consider  this  item  in  making  a  notebook. 
The  pencil-point  must  be  rubbed  upon  fine-  sandpaper  until  it  is  smooth 
and  conical.  The  beginner  should  draw  3  parallel  lines,  2  mm.  apart,  upon 
which  to  place  the  letters.  The  letters  may  be  placed  close  together  or 
farther  apart,  but  in  either  case  the  space  between  the  letters  must  be  kept 
the  same.  Larger  spaces  are  left  between  words.  Letters  may  be  straight 

ABCDEFGHIJKLMNOPQRSTUVWXYZ 
abcdefghijklmnopqrstuvwxyz 
0123456789 

FIG.  43. — Simplified  Gothic  or  "Shop  Skeleton"  Letters  and  Figures  Used  in  Label- 
ing Drawings. 

or  slanted  to  suit  the  individual  taste.  If  one  has  never  done  any  labeling, 
time  should  be  taken  to  practice  upon  a  piece  of  paper  before  finishing  a 
drawing.  Many  letters  like  b,  d,  and  g  contain  an  o  combined  with  a  straight 
line,  hence  it  is  necessary  to  learn  to  make  an  o  properly.  In  making  b,  d,  p, 
and  g,  be  careful  that  the  up  or  down  stroke  is  straight  and  joins  the  curve 
smoothly.  The  b,  d,  /,  h,  k,  and  I,  extend  above  the  other  letters  to  the 
height  of  capitals,  while  g,  j,  p,  q,  and  y  extend  just  as  far  below  the  line. 
The  t  falls  between  the  stem  letters  and  the  short  letters.  The  Gothic  style 
given  in  Fig.  43  is  easy  to  learn  and  will  answer  all  purposes. 

H.    PREFERABLE  MODES  OF  REPRESENTATION  FOR 
SPECIAL  COURSES 

General. — Methods  of  representation  vary  as  the  subject-matter 
in  each  instance  requires  different  handling.  -The  modes  here 
described  have  been  found  practical  in  different  biological  courses; 
however,  each  may  be  altered  to  suit  individual  requirements.  In 


Drawing  165 

elementary  courses  in  general  zoology  unshaded  ink-drawings  are 
best.  They  are  more  accurately  executed  by  elementary  students 
because  defects  are  so  obvious  that  they  do  not  pass  unnoticed; 
moreover,  the  student  exercises  more  care  in  making  the  drawing 
because  of  the  greater  difficulty  of  changing  it  after  it  is  once  drawn. 
Such  drawings  may  be  made  more  or  less  diagrammatic,  depending 
upon  the  ability  of  the  student  and  his  previous  training.  Students 
with  no  former  experience  in  representing  upon  paper  what  they  see 
need  not  be  discouraged,  for  clear-cut  outline  drawings  can  be  made 
by  any  one,  if  due  consideration  is  given  to  the  points  enumerated  under 
the  foregoing  paragraphs. 

Embryology. — The  first  step  in  embryological  drawing  is  a  care- 
ful outline  with  a  4H  pencil.  As  an  aid  in  drawing  complex  sections 
— a  cross-section  of  an  old  tadpole,  for  example — a  cover-glass 
ruled  into  squares  can  be  fastened  into  the  ocular.  The  drawing- 
paper  is  ruled  into  the  same  number  of  squares  as  the  cover-glass; 
each  square  as  many  times  the  size  of  one  square  of  the  cover-glass 
as  the  intended  magnification.  Parts  of  the  object  under  the 
squares  in  the  ocular  can  be  located  in  corresponding  squares  upon  the 
paper  (Isaacs,  Anatomical  Record,  IX,  711-13).  In  such  drawings 
cells  are  not  usually  indicated.  It  is  a  decided  advantage  to  have 
the  parts  colored,  especially  if  organs  from  the  same  germ  layer  are 
colored  alike.  If  three  colors  are  chosen,  one  for  each  of  the  germ 
layers,  ectoderm  (blue),  endoderm  (yellow),  and  mesoderm  (red), 
and  if  these  are  consistently  used  throughout  the  sections,  one  can 
see  at  a  glance  the  development  of  the  organs.  Crayon  pencils  work 
up  rapidly  and  give  good  results  if  the  color  is  put  on  lightly,  so 
that  in  the  finished  drawing  the  colors  blend.  To  spread  crayon 
evenly,  use  a  blunt,  rounded  point  and  make  long  strokes  with  the 
side  of  the  crayon.  For  example,  in  a  cross-section  of  a  frog  embryo, 
the  neural  tube  with  its  optic  cups  is  ectodermal  in  origin  and  the  blue 
color  immediately  indicates  its  relationship  to  the  ectoderm  which  is 
similarly  colored;  the  mesoblastic  somites  and  mesenchyme  are  red; 
the  lining  of  the  archenteron  yellow.  In  older  embryos  where  many 
parts  develop  from  the  mesoderm,  other  colors  may  be  used  for  special 
parts — as  brown  for  kidneys.  The  shade  of  red  may  likewise  be 


166  Animal  Micrology  - 

varied,  a  dark  red  being  used  for  blood  vessels,  obtained  by  putting 
on  a  heavy  layer  of  crayon ;  while  a  light  red  may  be  used  for  mesen- 
chyme.  By  this  method  a  student  has  constantly  before  him  the 
layers  from  which  various  organs  develop,  while  the  instructor  can 
immediately  see  that  the  student  has  or  has  not  a  clear  conception  of 
the  manner  in  which  the  organism  is  built  up.  Water  colors  can 
be  used  instead  of  crayons,  but  in  the  hands  of  most  students  they 
are  less  satisfactory.  Where  cellular  structure  is  put  in,  colored 
inks  may  be  employed.  However,  drawing  the  individual  cells  in  an 
embryo  is  too  time-consuming  a  process  for  ordinary  class  work  in 
embryology. 

Histology. — Histological  drawings  are  best  executed  in  pencil. 
Details  must  be  shown.  In  general,  these,  especially  nuclear  dif- 
ferences, can  be  put  in  by  stippling.  In  intercellular  matrices,  such 
as  connective  tissue,  the  texture  of  the  tissue  should  be  represented 
by  irregular  lines.  Light  lines  give  the  effect  of  fibers  and  fibrils  very 
well.  Where  different  tissues  of  an  entire  organ  are  to  be  distin- 
guished, the  whole  drawing  may  be  covered  with  a  light  ground- 
substance  of  blended  graphite  and  the  details  worked  up  with  pen 
and  ink.  This  combination  is  effective  and  has  the  added  advantage 
of  quick  execution. 

Cytology. — Cytology  requires  even  more  detail  than  histology. 
Stippling  or  wash  is  used  chiefly.  A  different  arrangement  of  the 
dots  gives  the  granular,  alveolar,  or  reticular  appearance  of  cyto- 
plasm. Solid  lines  should  be  avoided  as  much  as  possible;  fibers  can 
be  represented  by  dots  placed  close  together  in  a  linear  row.  Chromo- 
somes can  be  stippled,  made  solid,  or  blended  with  graphite  over 
their  entire  area.  Where  the  cell  cytoplasm  is  homogeneous,  a  light 
ground-coat  of  graphite  can  be  placed  over  the  entire  cell  and  the 
cell  parts  stippled  upon  this  with  pen  or  pencil.  A  wash  can  be 
substituted  for  the  graphite;  but  as  it  requires  more  careful  applica- 
tion it  is  not  recommended  for  class  work. 

Crayons  used  judiciously  in  both  histological  and  cytological 
drawings  are  often  effective.  Secretory  granules,  where  present,  can 
be  put  in  with  color  if  the  crayons  are  sharpened  to  a  fine  point. 
Ground-substance  of  tissues  can  be  depicted  with  crayon  and  the 


Drawing  167 

characteristic  features  of  the  tissue  added  with  pen  or  pencil.  Gran- 
ules of  white  blood  corpuscles  have  definite  color  reactions  upon  the 
basis  of  which  they  are  classified;  such  granules  should  therefore  be 
colored  in  a  drawing. 

m.     DRAWINGS  FOR  PUBLICATION 

To  make  illustrations  for  publication  in  a  book  or  scientific 
journal,  one  must  not  only  understand  form,  color,  perspective,  and 
composition,  but  also  know  something  about  the  science  of  repro- 
ducing drawings  in  printed  form.  Manuscripts  which  contain 
drawings  that  can  be  cheaply  reproduced  are  more  readily  accepted 
for  publication  than  those  which  require  expensive  plates.  Before 
undertaking  a  series  of  drawings  for  publication  make  a  careful 
study  of  similar  work  in  standard  journals,  particularly  in  the  journal 
in  which  you  expect  to  publish. 

Materials  for  Manuscript  Drawings. — In  general,  drawings  for 
publication  should  be  made  in  black  and  white,  because  this  style 
can  be  reproduced  most  cheaply  by  publishers.  For  working  with 
ink,  a  water-proof  India  ink,  such  as  Higgins',  is  best.  This  can 
be  applied  with  pen  or  brush.  Gillott's  penpoints  are  satisfactory. 
Inasmuch  as  each  person  not  only  handles  a  pen  differently,  but 
uses  different  degrees  of  pressure  in  working,  he  must  determine 
by  experience  the  number  of  the  pen  best  suited  to  his  needs.  For 
fine  linework  and  stippling,  the  writer  has  found  that  Gillott's 
lithographic  pen  No.  290  gives  the  best  results.  Fine  red-sable 
brushes  may  also  be  used  for  this  work,  although  for  line-process 
reproduction  (p.  168)  the  pen  drawing  is  likely  to  prove  more  suc- 
cessful. 

Drawings  should  be  made  upon  a  good  quality  of  paper.  Bristol 
board,  either  2-  or  4-ply,  or  Whatman's  hot-pressed  (smooth)  water- 
color  paper,  either  of  which  can  be  obtained  at  any  store  carrying  a 
complete  line  of  stationery  materials,  can  be  used.  Whatman's 
paper  is  of  the  same  texture  throughout;  moreover,  it  can  be  used 
for  ink,  wash,  or  pencil-work.  Anvil  drawing-paper,  No.  105,  is 
excellent  for  large  illustrations  or  charts.  This  is  a  cloth-backed 
paper  which  can  be  rolled  without  injury.  A  disadvantage  is  the 


168  Animal  Micrology 

large  size  and  expense  of  a  roll,  which  contains  10  yards  of  paper 
36  inches  wide.  It  is  put  out  by  the  Keuffel  &  Esser  Co.,  of  New 
York.  Cloth-backed  papers  can  be  obtained  from  other  firms.  A 
stipple-board  (Ross  board),  manufactured  by  the  Charles  J.  Ross 
Co.,  consists  of  a  chalk  surface  upon  a  paper  back.  The  advantage 
of  this  paper  is  the  rapidity  with  which  drawings  can  be  made.  A 
stipple  effect  is  obtained  simply  by  rubbing  the  flat  side  of  a  pencil- 
point  back  and  forth  over  the  chalk  surface.  Different  stipple 
effects  are  obtained  by  using  different  grades  of  the  paper.  For  all 
general  uses  No.  8  is  best. 

Water  colors  may  be  procured  in  a  variety  of  forms  and  makes 
for  wash-drawings.  On  account  of  the  cost  of  reproduction,  however, 
in  papers  which  are  to  be  published  colored  drawings  should  be 
avoided  wherever  possible.  Winsor  &  Newton's  Ivory  Black  or 
Charcoal  Grey,  which  comes  in  a  cake,  is  best  for  black-and-white 
wash-drawings.  Never  use  blue  in  a  drawing  to  be  photographed, 
as  it  does  not  take.  Remember  also  that  yellow  and  brown  appear  as 
black.  Likewise  keep  in  mind  that  the  results  are  in  no  appreciable 
degree  dependent  upon  the  number  or  kind  of  tools  used,  but  upon 
the  skill  shown  in  execution. 

Camera  Lucida. — As  an  aid  in  making  drawings  of  microscopic 
objects,  an  instrument  known  as  the  camera  lucida  is  often  employed. 
With  such  an  instrument  the  image  of  an  object  under  the  microscope 
can  be  projected  upon  the  drawing-paper  (see  p.  189). 

Reduction  of  Drawings. — In  making  drawings  for  publication, 
it  is  advisable  to  make  them  larger  than  they  will  appear  in  the 
finished  cut,  as  in  the  reduction  many  irregularities  are  lessened. 
But  under  no  circumstances  make  a  crude  drawing  with  the  idea 
that  in  the  print  it  will  appear  perfect,  for  while  reduction  minimizes, 
it  does  not  obliterate  defects.  The  original  drawing  should  not 
ordinarily  be  more  than  twice  the  size  of  the  intended  cut,  while  a 
reduction  of  one-fourth  or  one-third  will  probably  give  a  better 
result.  If  one  is  not  careful  about  the  spacing  of  lines  and  dots  used 
in  the  drawing,  reduction  tends  to  make  them  run  together. 

Line-Process. — Line-process  is  not  only  the  cheapest  form  of 
reproduction  used  by  journals,  but  likewise  the  most  accurate. 


Drawing  169 

Prints  reproduced  in  this  manner  contain  contrast  not  gained  by  any 
other  method.  The  drawing  is  photographed  directly  upon  a  zinc 
plate.  Ink  is  then  applied  to  this  plate  with  an  ordinary  ink-roller. 
The  sticky  ink  adheres  only  to  the  lines  of  the  photograph.  The 
inked  plate  is  now  placed  in  a  bath  of  acid  which  eats  away  the 
uninked  portions.  The  metal  plate,  containing  the  design  in  relief, 
mounted  upon  wood,  type  high,  is  set  up  and  printed  with  the  type 
upon  text-paper  if  the  details  are  not  too  fine  and  if  no  colors  are 
required. 

Lines. — For  this  type  of  reproduction  use  ink  as  the  working 
medium  and  apply  it  with  pen  upon  smooth  white  bristol  board 
or  water-color  paper.  Lines  should  not  be  extremely  fine,  and  care 
should  be  taken  that  they  are  far  enough  apart  to  reproduce  well. 
Likewise,  cross-hatching,  if  employed,  must  be  coarse  enough  to 
stand  the  reduction,  otherwise  it  will  appear  as  a  solid  black  mass. 
While  ink  thinned  to  a  gray  gives  a  difference  in  tone  in  the  original 
drawing,  it  must  be  borne  in  mind  that  this  behaves  erratically  in 
reproduction.  Gray  lines  often  appear  in  the  print  as  broken 
black  lines,  or  they  may  be  entirely  lost.  To  avoid  graying  black 
lines  the  pen  must  be  wiped  frequently  and  filled  with  a  fresh 
supply  of  ink.  To  get  effects  in  tone  vary  either  the  thickness 
of  the  lines  or  the  distance  between  the  lines.  Lines  in  the  fore- 
ground should  be  farther  apart  and  heavier  than  those  in  the 
background. 

Dots. — If  drawings  are  stippled,  the  dots  should  be  made  with  the 
amount  of  reduction  in  mind.  Too  fine  stippling  necessitates 
etching  upon  copper,  a  tedious  process,  which  doubles  the  cost  of 
production.  To  secure  degrees  of  light  and  shade,  vary  the  distance 
between  the  dots,  not  the  size .  of  the  dots.  Lithographic  'crayon 
used  upon  stipple-board  (see  p.  168)  will  reproduce  by  line-process  if 
the  drawing  is  coarse  enough  to  stand  a  reduction  to  one-half  its 
size. 

Graphs. — Graphs  for  line  reproduction  should  be  made  upon 
co-ordinate  paper  in  which  the  lines  are  blue.  As  blue  does  not 
photograph,  the  co-ordinates,  both  perpendicular  and  horizontal, 
which  are  to  appear  in  the  cut  must  be  inked. 


170  Animal  Micrology     , 

Color. — A  separate  block  and  printing  are  required  for  each  color 
used  in  line  engraving.  In  a  drawing  in  which  several  colors  are  to 
appear,  the  original  drawing  in  black  and  white  must  be  accompanied 
by  a  colored  sketch.  From  a  process-plate  of  the  black-and-white 
drawing  a  print  is  taken.  For  each  color  used  in  the  sketch,  the 
printer  then  makes  separate  blocks  which  are  printed  one  over  the 
other  upon  the  print  from  the  first  plate.  Avoid  fine  gradations  in 
color,  as  thereby  the  expense  of  reproduction  is  increased. 

Half-Tone. — Shaded  and  wash-drawings  and  photographs  are 
ordinarily  reproduced  by  the  half-tone  process.  A  screen  containing 
133  to  175  meshes  per  square  inch  is  placed  between  the  camera  and 
the  drawing  when  the  latter  is  photographed.  This  is  done  to  break 
up  the  flat  tones  into  dots,  otherwise  they  would  print  as  black 
masses.  The  photograph  thus  made  is  transferred  to  a  zinc  or 
copper  plate  and  etched  in  the  same  way  as  a  line-process  plate. 
Where  the  details  of  the  picture  are  not  too  delicate,  the  print  can 
be  made  upon  text-paper.  For  proper  reproduction,  drawings  which 
require  a  fine  screen  must  be  carefully  printed  upon  special 
coated  paper. 

It  must  be  remembered  that  the  screen  on  account  of  its  fine 
dots  introduces  shadow  into  the  lighter  parts  of  the  picture;  while 
the  white  spaces  between  the  dots  render  the  darker  areas  lighter. 
These  changes  demand  stronger  contrasts  in  the  original  drawings 
and  photographs  which  are  to  be  reproduced  by  half-tone  than  those 
desired  in  the  print. 

Wash-drawings. — Wash-drawings  (p.  162)  for  reproduction  should 
not  contain  solid  black  lines,  as  the  screen  breaks  up  the  lines  into 
dots.  Be  careful  that  no  pencil  lines  remain  in  the  drawing  and 
avoid  smudges  and  finger-marks.  Jf  clear  white  spaces  are  desired 
in  the  print,  the  shadows  produced  by  the  screen  in  these  places  must 
be  tooled  out  of  the  plate  to  give  the  required  effect — a  process  which 
is  slow  and  expensive. 

Combination  with  wash. — Drawings  can  be  made  in  which  certain 
parts  are  put  in  with  pen  or  brush  lines  upon  a  wash-background. 
This  makes  a  satisfactory  combination  for  half-tone  prints.  On  the 
other  hand,  fine  pencil  striae  upon  a  wash-background  are  usually  lost 


Drawing  171 

in  reproduction,  since  they  offer  insufficient  contrast  with  the  back- 
ground. 

Pencil. — Pencil-work  does  not  reproduce  satisfactorily  by  half- 
tone. Where  blended  graphite  is  used  for  a  background  instead  of 
wash,  a  reproduction  can  be  made  if  the  print  is  made  upon  good 
coated  paper.  However,  the  results  are  so  inferior  to  the  originals 
that  it  pays  to  take  the  time  to  learn  how  to  make  good  wash- 
drawings. 

Color. — Color  can  be  applied  upon  half-tone  prints,  although  a 
separate  block  and  printing  are  necessary  for  each  color.  Plates 
are  sometimes  made  as  described  under  "Line-Process,"  and  these 
printed  in  color  upon  a  half-tone  background  (see  p.  170). 

Photographs. — Photographs  are  usually  reproduced  by  half- 
tones. A  hard-finish,  glossy  paper  which  brings  out  strong  contrast 
between  the  blacks  and  whites  of  the  picture  is  best  for  prints. 
Azo  hard  X,  glossy  white  Velox  paper  made  by  the  Eastman  Kodak 
Co. ,  and  solio  paper  of  a  brownish  tinge  reproduce  well.  Photographs 
will  stand  some  reduction,  but  on  the  whole  it  is  better  to  have 
prints  the  same  size  as  the  intended  cut.  Prints  should  be  squee- 
geed. This  is  done  with  a  roller  which,  passed  over  the  wet  print, 
removes  the  moisture  and  gives  a  hard  finish  to  the  gelatin  film 
when  it  dries. 

Lithography. — Lithography  is  the  most  expensive  form  of  repro- 
duction. While  line-process  and  half-tone  plates  are  made  by 
mechanical  means,  lithographic  work  is  all  done  by  hand.  A  picture 
to  be  reproduced  by  this  method  must  be  transferred  to  a  stone  and 
the  parts  cut  in  with  a  graver  in  the  hand  of  an  expert.  Undoubtedly 
it  is  the  most  artistic  form  of  reproduction,  but  most  journals  will 
not  accept  drawings  wrhich  have  to  be  engraved  unless  the  author  or 
artist  pays  the  extra  cost  of  reproduction.  Intricate  drawings  of 
many  colors  which  cannot  be  reproduced  by  line-process  or  half-tone 
come  out  well  by  this  method,  if  cost  is  not  an  item  to  be  considered. 
Likewise  stippled  pencil  drawings  make  excellent  illustrations  with 
this  sort  of  reproduction.  Heliotype  (gelatin  plate)  is  a  some- 
what less  expensive  method  for  reproducing  colored  drawings  and 
photographs. 


172  Animal  Micrology 

Arrangement  of  Drawings  for  Reductions. — Line-drawings  where 
not  used  as  text-figures  should  be  arranged  in  the  form  of  plates. 
Half-tones  are  usually  so  arranged,  since  they  generally  require 
a  special  coated  paper.  To  arrange  drawings  in  the  form  of  a  plate, 
one  must  know  the  exact  amount  they  are  to  be  reduced  when 
printed.  If  one-half,  for  instance,  they  must  be  arranged  as  a  plate 
twice  the  size  of  the  journal  page.  The  drawings  can  either  be  made 
directly  upon  bristol  board  in  the  order  in  which  they  are  to  be 
printed,  or  they  may  be  pasted  upon  the  board  in  such  order.  The 
latter  method  is  usually  practiced.  Due  allowance  must  be  made 
for  lettering  and  the  margins  of  the  page. 

Lettering. — All  the  original  drawings  should  be  so  lettered  that  the 
letters  will  be  of  the  proper  size  when  reduced.  Letters  can  either 
be  pasted  on  or  printed  by  hand  (p.  164).  A  drawing  presents  a 
neater  appearance  if  the  lettering  is  parallel  to  the  base  line.  A  cut 
is  more  legible  if,  instead  of  abbreviations,  the  names  printed  in  full 
are  connected  to  the  proper  part  of  the  drawing  by  leaders.  Gummed 
sheets  containing  letters,  numerals,  and  such  words  as  "Plate"  and 
"Fig."  in  several  sizes  can  be  bought  for  use  in  this  work.  Likewise 
publishers  of  some  journals  will  print  letters  and  words  which  can 
be  pasted  on. 

For  a  good  chapter  on  laboratory  drawing  see  A  Laboratory 
Guide  for  Histology,  by  Irving  Hardesty  (P.  Blakiston's  Son  &  Co., 
Philadelphia) . 


APPENDIXES 


APPENDIX  A 


THE  MICROSCOPE  AND  ITS  OPTICAL  PRINCIPLES 

For  an  understanding  of  the  optical  principles  involved  in  micros- 
copy, four  things  must  be  borne  in  mind  with  regard  to  a  ray  of 
ordinary  daylight: 

1.  It  has  an  appreciable  breadth. 

2.  It  travels  in  a  straight  line  in  a  homogeneous  medium. 

3.  It  is  bent  (refracted)  in  passing  obliquely  from  one  medium 
into  another  of  different  density. 

4.  It   is   in  reality   a  composite  of  a 
number  of  different  colored  rays,  ranging 
from  violet  to  red,  and  each  of  these  has  a 
different  refrangibility. 

The  amount  of  refraction  undergone  by 
light  in  a  given  case  depends  upon  the 
difference  in  density  of  the  two  media  which 
the  light  traverses.  Thus,  glass  is  denser 
than  air;  hence,  in  passing  from  air 
obliquely  through  a  glass  plate  (Fig.  44),  a  ray  of  light  A  B  would 
be  bent  out  of  its  original  course.  On  reaching  the  air  again,  how- 
ever, it  would  resume  its  original  direction,  although  it  .would  be 
displaced  to  an  amount  equal  to  the  distance  between  A  and  A'. 

It  is  on  account  of  such  displacement 
that  an  object  in  water,  for  example, 
appears  to  be  at  a  different  point  from 
where  it  really  is. 

On  the  other  hand,  after  traversing 
a  prism,  a  ray  does  not  resume  its 
former  direction,  but  takes  a  new 
course  upon  leaving  as  well  as  upon  entering  the  prism  (Fig.  45). 
This  new  direction  is  always  toward  the  base  of  the  prism,  and  the 
amount  of  deviation  depends  upon  the  shape  and  density  of  the 

175 


FIG.  44 


FIG.  45 


176 


Animal  Micrology 


prism.  If  the  base  is  down,  then  the  ray  is  bent  downward;  if  the 
apex  is  down,  the  ray  still  deviates  toward  the  base,  that  is,  it  is 
bent  upward. 

Lenses. — Each  of  the  two  principal  forms  of  lenses  is  in  effect 
practically  two  prisms,  (1)  with  the  bases  placed  together  (Fig.  46,  a, 

convex  lens),  or  (2)  with  the  apexes  together 

(Fig.  46,  b,  concave  lens). 

In  the  convex  lens,  since  rays  of  light  are 

refracted  toward  the  bases  of  the  respective 

prisms,  they  will  converge;   in  the  concave 

lens,  for  the  same  reason,  they  will  diverge. 

The  terms  converging  lens  and  diverging  lens, 
b          therefore,  are  used  frequently  as  synonymous 

with  the  terms  "convex  lens"  and  "concave 
lens."  All  lenses  are  modifications  or  combinations  of  these  two 
types. 

If  parallel  rays  of  light  pass  through  a  convex  lens  (Fig.  47)  they 
are  so  refracted  as  to  meet  in  one  point  F,  which  is  termed,  in  conse- 
quence, the  focal  point  or  principal  focus.  If,  on  the  other  hand,  the 
source  of  light  be  placed  at 
the  focal  point,  then,  after 
traversing  the  lens,  the  rays 
of  light  will  emerge  parallel. 
If  parallel  rays  of  light  came 
from  the  opposite  side  of  the 
lens,  manifestly  there  would 
be  a  second  focal  point  at  F' . 
The  two  principal  foci  are 
termed  conjugate  foci,  and  will 
be  equidistant  from  the  center 
of  the  lens  when  both  sides  of  the  lens  have  equal  curvature 
The  ray  which  passes  through  the  center  of  the  lens  (Fig.  47,  c) 
and  the  focal  point  traverses  what  is  termed  the  principal  axis  of 
the  lens.  The  optical  center  of  the  lens  is  a  point  on  the  principal 
axis  through  which  rays  pass  without  angular  deviation.  It  may 
be  within  or  outside  the  lens,  depending  upon  the  form  of  the  latter. 


FIG.  47 


The  Microscope  and  Its  Optical  Principles 


17? 


FIG.  48 


Any  line  (ed),  other  than  the  principal  axis,  which  passes  through 
the  optical  center  of  the  lens  is  termed  a  secondary  axis. 

In  the  case  of  a  concave  lens,  parallel  rays  will  be  caused  to 
diverge  (Fig.  48)  and  the  principal  focus,  F,  of  the  lens  is  determined 
by  the  extension  of  the  divergent  rays  till  they  meet  at  a  point  which 
lies  on  the  same  side  of  the 
lens  as  the  source  of  light. 

Such  a  point  has  no  actual     

existence,  and  is  known, 
consequently,  as  a  virtual 

focus.    The  focus  of  a  con-     r  ^ 

vex  lens,  on  the  other  hand, 
is  real,  and  may  be  deter- 
mined readily  by  allowing  

the  sun's  rays,  which  are 

practically  parallel,  to  pass 

through  it  on  to  a  screen. 

By  moving  the  lens  backward  and  forward,  the  spot  of  projected 

light  varies  in  size  and  brightness.     When  smallest  and  brightest 

the  spot  is  at  the  focal  point  of  the  lens. 

Images. — In  Fig.  49  the  object,  represented  by  an  arrow,  lies 
beyond  the  principal  focus  of  a  convex  lens  as  in  a  photographic 
camera,  for  example,  or  the  objective  of  a  compound  microscope. 

Light  rays  pass  out  in  all 
directions  from  any  lumi- 
nous point.  Hence,  one 
ray  from  any  point  on 
the  arrow,  the  tip,  for 
instance,  will  pass  through 
the  focal  point,  F,  and 

one  will  pass  through  the  optical  center  of  the  lens.  From  what 
was  determined  above,  manifestly  the  ray  through  F  will  emerge  as 
one  of  the  parallel  rays  upon  leaving  the  lens,  and  the  one  through 
the  optical  center  of  the  lens,  since  it  traverses  a  secondary  axis, 
will  not  be  refracted,  hence  the  two  rays  must  cross.  Their  point 
of  intersection  is  the  point  at  which  the  image  of  the  arrow-tip  will 


178 


Animal  Micrology 


be  formed.  The  same  fact  may  be  determined,  likewise,  for  any 
other  point  of  the  arrow,  for  example,  the  opposite  end.  Thus  the 
distance  from  the  lens  at  which  the  image  is  formed  may  readily  be 
determined.  In  focusing  a  photographic  camera,  for  example,  the 
image  comes  sharply  into  view  on  the  ground-glass  plate  at  the 
back  of  the  camera  when  the  plate  is  brought  into  the  plane  in 
which  these  rays  through  the  focus  and  the  optical  center  intersect 
beyond  the  lens.  It  will  be  observed  from  the  figure  that  the  image 
is  reversed.  The  size  of  the  image  diminishes  as  the  object  lies 
farther  beyond  F. 

In  case  the  object  lies  between  the  lens  and  the  principal  focus, 
as  in  Fig.  50,  parallel  rays  from  the  object  would  converge  to  meet 
at  the  conjugate  focus  Ff ',  and  an  eye  at  this  point  would  see  the 

image  projected  and  enlarged 
without  being  reversed.  The 
plane  in  which  the  image  is 
formed  is  determined  by  finding 
the  points  of  intersection  of  the 
secondary  axes  through  points  of 
the  object  with  the  imaginary 
elongation  of  the  refracted  rays 
as  shown  in  the  figure.  The 
image  is  magnified  because  the  observer  judges  of  the  size  of  an 
object  by  the  visual  angle  which  it  subtends.  The  greater  the  con- 
vexity of  the  lens  the  shorter  the  focus,  and  also,  since  the  rays  are 
bent  more,  the  greater  the  magnification. 

The  Simple  Microscope. — The  simple  microscope  (the  ordi- 
nary so-called  magnifier,  etc.)  operates  upon  this  principle;  the 
image  of  an  object  is  projected  and  enlarged  but  not  inverted 
(Fig.  50). 

The  question  arises  as  to  why  there  is  a  best  distance  to  hold  the 
simple  microscope  from  an  object.  Why  will  not  any  point  answer 
so  long  as  it  is  within  the  focal  point?  As  a  matter  of  fact,  the  object 
may  be  placed  at  any  point  within  the  focus,  and  it  will  be  found  that 
the  nearer  it  is  brought  to  the  lens  the  less  it  is  magnified.  There  is 
one  most  favorable  point  for  observation,  however,  which  is  neither 


PIG.  50 


The  Microscope  and  Its  Optical  Principles 


179 


at  the  point  of  highest  nor  of  lowest  magnification,  but  an  intermediate 
point,  where  the  lens  is  freest  from  chromatic  and  spherical  aberra- 
tions. 

In  reality  the  eye  forms  an  integral  part  of  the  optical  arrange- 
ment when  the  microscope  is  being  used,  but  in  our  elementary 
exposition  of  the  subject  it  is  disregarded. 

The  Compound  Microscope. — The  general  principle  of  the  com- 
pound microscope  is  represented  in  Figs.  51  and  58.  The  object  ab 
(Fig.  51)  lies  beyond  the  principal  focus  of  the  first  lens  or  objective 


FIG.  51 

(really  a  system  of  lenses),  hence  the  image  AB  is  reversed.  This 
image,  in  turn,  is  viewed  through  a  lens,  the  eyepiece  or  ocular  situated 
nearer  the  eye  of  the  observer.  The  ocular  acts  as  a  simple  magnifier, 
projecting  and  enlarging  the  image  but  not  reversing  it  again.  As  a 
matter  of  fact,  the  ordinary  ocular  of  a  compound  microscope 
cannot  be  taken  from  the  instrument  and  used  as  a  simple  magnifier 
because  it  is  made  of  two  planoconvex  lenses  which  are  so  adjusted 
that  the  image  from  the  objective  of  the  compound  microscope  is  not 
brought  to  focus  until  it  has  traversed  the  larger  or  field-lens  of  the 
eyepiece  (Figs.  55,  58).  The  image  is  really  examined,  therefore,  at 
a  point  between  the  two  lenses  of  the  eyepiece.  Such  an  eyepiece  is 


180  Animal  Micrology 

termed  a  negative  eyepiece  or  ocular  and  is  widely  used  today  for 
microscopical  work.  The  commonest  form,  the  Huygenian,  is  an 
adaptation  of  an  ocular  designed  originally  by  Huygens  for  the 
telescope.  By  contracting  the  area  of  the  real  image,  the  field- 
lens  of  a  negative  ocular  not  only  brightens  the  image  but  also 


i-Inch  Objective  J-Inch  Objective          T\-Inch  Oil-Immersion  Objective 

FIG.  52. — Lens  Systems  of  Various  Objectives 
Bausch  &  Lomb  f-inch,  i-inch,  and  T\  -inch  oil-immersion  objectives,  respectively 

increases  the  size  of  the  field  that  can  be  examined.  It  is  usually 
also  designed,  in  conjunction  with  the  eye-lens,  to  help  render  the 
image  achromatic. 

Positive  eyepieces  are  also  made.     An  inverted  image  of  the 
object  is  formed  below  the  system  of  ocular  lenses.     Such  an  ocular 

operates  as  a  simple 
microscope. 

A  good  objective  is 
made  up  of  from  two  to 
five   systems   of    lenses, 
as  shown  in  Fig.  52.     A 
PIG  53  single  system  in  turn  may 

be  a  doublet  (Fig.  57)  or 

a  triplet,  each  made  of  different  kinds  and  shapes  of  glass.  A 
good  objective  is  a  very  delicate  piece  of  apparatus  and  must  be 
handled  with  great  care.  Each  component  is  very  accurately  ground 
and  the  systems  distanced  with  extreme  precision  in  order  to  get 
a  clear  image.  If  not  already  familiar  with  the  parts  of  the  com- 


The  Microscope  and  Its  Optical  Principles 


181 


pound  microscope,  the  student  should  study  Figs.  54,  55,  and  58, 
with  a  microscope  before  him. 

DEFECTS  IN  THE  IMAGE 

Spherical  Aberration. — A  simple  convex  lens,  unless  corrected, 
will  not  give  a  sharply  defined  image  because  it  does  not  refract 


berHead  or 
Fine  Adjustment: 

Arm. 


-CUP..N 

,  TV; sin  WasHer. 


_ — Horse.  Shoe  Base. 


FIG.  54. — A  Compound  Microscope  with  Parts  Named 

to  the  same  degree  all  rays  passing  through  it.  Those  which  traverse 
its  edges  are  brought  to  a  focus  nearer  the  lens  (Fig.  53).  This 
results  not  only  in  an  indistinct  image  but  in  a  distortion  of  shape  as 


182 


Animal  Micrology 


Body  Tube 


") 

I  of  the  Ocular 
•Diaphragm/ 
-•Field  lens/ 
— -DrawTube 


.  ,D  raw  Tube  Diaphra  gm 
with  Society  Screw 


—.Society  Scraw 


-Baonlens      v  ,  ,h 

iddlelens     I  J 
.front  lens     \Obje.cUve 
'..WorkingDbtance/ 


.Slide' 


PIG.  55. — Sectional  View  of  Microscope  Tube 
Including  Ocular  and  Objective. 


well.  Straight  lines,  for 
example,  appear  curved,  and 
when  the  parts  of  the  object 
are  in  focus  in  the  center  of 
the  field,  those  nearer  the 
margin  are  hazy  and  indis- 
tinct. This  defect  is  greatest 
in  strongly  curved  lenses,  that 
is,  since  magnification  in- 
creases with  increased 
curvature,  in  high  powers. 
Spherical  aberration  is 
corrected  by  one  or  more  of 
the  following  processes: 

1.  Cutting  off  the  margi- 
nal rays. 

2.  Changing  the  shape  of 
the  surface  of  the  lens. 

3.  Combining     several 
lenses  equivalent  to  a  single 
lens. 

Chromatic  Aberration. — • 
As  with  a  prism,  ordinary 
light  in  passing  through  a  lens 
is  broken  up  into  its  compo- 
nent colors.  This  process  is 
Since  the  colors  are  not  all  bent  to 


technically  termed  dispersion. 

the  same  extent,  the  result  is  that 

each  color  has  a  different  focus;  the 

ones  which  are  bent  most  (violet 

rays)  come  to  a  focus  nearest  the 

lens,   and    those   which   are   least 

affected  (red  rays)  meet  at  a  point 

farther  away   (Fig.   56).     This 

failure  of  the  color  rays  to  meet  in 

one  focal  point  is  termed  chromatic  aberration,  and  if  unconnected 


The  Microscope  and  Its  Optical  Principles  183 

causes  the  image  of  an  object  viewed  through  such  a  lens  to  be 
bordered  by  a  colored  halo. 

The  defect  is  corrected  by  properly  combining  glasses  of  different 
dispersive  powers  but  of  kindred  refractive  powers.  Flint  glass 
(silicate  of  potassium  and  lead),  for  example,  has  a  dispersive  power 
equal  to  about  twice  that  of  crown  glass  (silicate  of  potassium  and 
lime),  although  their  refractive  powers  are  nearly  the  same. 
By  combining  a  biconvex  lens  of  crown  glass  with  a  con- 
cave lens  of  flint  glass  so  constructed  that  its  dispersive 
power  will  just  equal  that  of  the  crown  glass  (Fig.  57),  the 
error  may  in  large  measure  be  corrected.  Such  an  arrange-  piQ  5? 
ment  does  not  interfere  seriously  with  the  refractive  powers 
of  the  lens  so  constructed.  Unfortunately  no  two  kinds  of  glass 
have  been  found  which  have  proportional  dispersive  powers  for  all 
colors,  so  that  in  the  ordinary  achromatic  objective  only  two  of  the 
different  colors  of  the  spectrum  have  been  accurately  corrected  and 
brought  to  one  focus.  The  colors  left  outstanding  form  the  defect 
known  as  a  secondary  spectrum.  In  the  apochromatic  objectives 
(p.  188)  three  rays  are  brought  to  one  focus,  leaving  only  a  slight 
tertiary  spectrum. 

NOMENCLATURE  OR  RATING  OF  OBJECTIVES 
AND  OCULARS 

Oculars. — Different  makers,  unfortunately,  use  different  systems 
in  marking  their  lenses  to  indicate  relative  powers  of  magnification. 
In  the  case  of  lettering  the  system  is  wholly  arbitrary;  the  only  rule 
is  that  the  nearer  to  A  the  letter  is  the  lower  the  magnification. 
When  the  objective  bears  a  figure  it  is  usually  indicative  of  the  magni- 
fying power  of  the  part  marked.  Thus  a  TV-inch  objective  magnifies 
approximately  120  diameters;  a  f-inch,  80  diameters;  a  J-inch,  20 
diameters;  a  1-inch,  10  diameters;  a  2-inch,  5  diameters;  and  so  on. 
This  means  that  an  objective  which  forms  an  image  10  times  the  real 
diameter  of  the  object  itself,  on  a  screen  placed  10  inches  (the  con- 
ventional distance  of  vision)  from  its  back  lens,  is  rated  as  a  1-inch 
objective.  If  it  formed  an  image  only  5  times  the  real  diameter  of  the 
object  it  would  be  a  2-inch  objective;  if  30  times,  a  f-inch  objective, 


184 


Animal  Microtogy 


FIG.  58. — Diagram  Showing  Path  of  Light  Rays  through  the  Compound  Micro- 
scope Together  with  Images  (from  Bausch  and  Lomb  catalogue). 

Fi,  upper  focal  plane  of  objective;  Fz,  lower  focal  plane  of  eyepiece;  A,  optical  tube 
length  =  distance  between  Fi  and  Fz;  Oi,  object;  Oz,  real  image  in  Fz,  transposed  by  the 
collective  lens,  to  Oi,  real  image  in  eyepiece  diaphragm;  Oz,  virtual  image  formed  at 
the  projection  distance  C.  250  mm.  from  EP,  eyepoint;  CD,  condenser  diaphragm; 
L,  mechanical  tube  length  (160  .mm.;:  1,  2,  3,  three  pencils  of  parallel  light  coming  from 
different  points  of  a  distant  illuminant,  for  instance,  a  white  cloud,  which  illuminate 
three  different  points  of  the  object. 


The  Microscope  and  Its  Optical  Principles  185 


PIG.  59. — The  Spencer  No.  15  Compound  Microscope 


186 


Animal  Micrologtf 


PIG.  60. — The  Bausch  &  Lomb  CAS  Compound  Microscope 


The  Microscope  and  Its  Optical  Principles  187 

and  so  on.     Such  magnification  is  termed  the  initial  magnifying 
power  of  the  objective. 

The  objectives  of  most  manufacturers  are  now  rated  in  milli- 
meters and  the  conventional  distance  of  vision  taken  as  250  mm. 
An  objective  of  3  mm.  focus,  therefore,  yields  an  initial  magnification 
of  83.3  diameters  (JX 250  =83. 3).  Compensating  oculars  (see 
below)  bear  numbers  which  indicate  the  number  of  times  the  eyepiece, 
when  used  at  a  -given  tube-length,  increases  the  initial  magnification. 
Ocular  12,  for  example,  with  a  3-mm.  objective  would  yield  a  magni- 
fication of  83.3X12=1,000  diameters,  with  a  standard  length  of 
tube.  Unfortunately  this  simple  system  does  not  apply  to  most 
ordinary  oculars,  which  are  more  or  less  arbitrarily  lettered  or 
numbered. 

SOME  COMMON  MICROSCOPICAL  TERMS  AND  APPLIANCES 

(Alphabetically  Arranged) 

Achromatic  Objective. — An  objective  corrected  for  chromatic  aberra- 
tion (p.  182).  The  correction  is  not  absolute. 

Achromatism. — Freedom  from  chromatic  aberration. 

Angular  Aperture. — The  angle  (measured  in  degrees)  formed  at  the 
point  of  focus  (F,  Fig.  62)  by  the  outermost  rays  (aF,  bF)  which  traverse 
the  objective  to  form  an  image.  This  angle  is  an  important  consideration 
because  on  it  depends  in  large  measure  the  defining  or  resolving  power  of 
the  objective.  It  is  evident  that  the  larger  the  angle  is  the  greater  the  num- 
ber of  rays  of  light  that  will  be  admitted  from  an  object.  Thus  the  object 
will  be  better  defined  to  the  eye.  In  low  powers  the  angle  may  be  very  wide, 
in  high  powers  it  must  necessarily  be  small.  Two  objectives,  even  though 
they  may  possess  different  powers  of  magnification,  will  have  the  same  bril- 
liancy if  they  are  of  the  same  angular  aperture;  on  the  other  hand,  if  they 
have  the  same  magnifying  power,  but  differ  in  angular  aperture,  the  bril- 
liancy is  reduced  in  the  one  of  smaller  angle.  In  immersion  lenses  the  liquid 
used  between  the  lens  and  the  object,  by  reducing  refraction,  has  the  effect 
of  increasing  the  angle  of  aperture.  See  "Immersion  Objective,"  also 
"Numerical  Aperture"  (pp.  195,  199). 

Apertometer. — An  instrument  for  measuring  both  the  angular  and  the 
numerical  aperture  of  objectives.  It  is  fitted  to  the  stage  of  the  microscope. 

Aplanatism — Freedom  from  spherical  aberration  (p.  181).  The  result 
is  a  flat  field  as  viewed  through  the  microscope.  Aplanatic  lenses  are  usually 
also  achromatic. 


188 


Animal  Micrology 


Apochromatic  Objective. — An  improved  form  of  objective  which  is  more 
exactly  achromatic  than  the  ordinary  objective  because  it  is  corrected  for 
•rays  of  three  colors  instead  of  two,  and  this  correction  is  equally  good  in 
all  parts  of  the  field.  In  the  ordinary  achromatic  objective  after  correc- 
tion there  is  a  residue  of  color  which  is  known  as  the  secondary  spectrum. 
In  the  apochromatic  lenses  correction  is  made  for  a  third  color,  and  usually 


FIG.  61. — The  Bausch  &  Lomb  KA  Binocular  Dissecting  Microscope,  Greenough 
Type. 

only  a  slight  tertiary  spectrum  is  left  uncorrected.  Spherical  aberration 
is  also  more  fully  corrected.  Furthermore,  in  these  objectives  the  foci 
of  the  optical  and  the  chemical  rays  are  identical,  hence  the  lenses  are  well 
adapted  to  photography.  In  the  glasses  of  the  apochromatics,  silicon  is 
replaced  by  boron  in  the  flint  series  and  by  phosphorus  in  the  crown  series. 
Fluorite  was  used  in  conjunction  with  the  glasses  in  the  earlier  forms  of 
apochromatic  lenses  with  the  result  that  the  lenses  frequently  deteriorated 


The  Microscope  and  Its  Optical  Principles  189 

in  warm,  moist  climates.  Several  makers  are  now  able  to  construct  apochro- 
matic  objectives  without  the  use  of  fluorite.  Both  dry  and  immersion 
apochromatics  are  made. 

Binocular  Microscope. — A  microscope  adapted  to  vision  with  both  eyes 
at  once.  One  of  the  most  important  advances  in  microscopy  during  the 
past  ten  years  has  been  the  development  of  binocular  microscopes  with 
erecting  prisms  which  enable  one  to  carry  on  dissections,  to  study  thick 
injected  preparations,  and  to  perform  other  manipulations  under  higher 
power  and  otherwise  more  advantageously  than  formerly.  In  general,  they 
consist  (Fig.  61)  of  two  optically  distinct  tubes  so  combined  that  the  objec- 
tives focus  on  the  same  point  from  different  angles.  A  magnified,  stere- 
oscopic vision  is  thereby  provided,  so  that  objects  which  have  depth  stand 
out  in  pronounced  relief.  The  upper  parts  of  the  tubes  may  be  rotated  so 
as  to  adjust  the  eye-points  of  the  oculars  to  the  width  between  the  pupils 
of  the  observer's  eyes.  If,  upon  closing  one  eye  and  then  the  other,  an 
image  is  not  seen  by  each  eye  without  moving  the  head,  the  eye-points  are 
too  close  together  or  too  far  apart.  The  oculars  should  be 
separated  or  approximated  accordingly.  When  they  are  s — \ 

correctly  adjusted  one  should  get  a  distinct,  stereoscopic       at J^ 

appearance.    Other  adjustments  are  provided  to  compensate          \       I 
for  differences  of  focus  in  the  right  and  the  left  eye.  \/ 

A  very  simple  form  of  binocular  magnifier,  known  as 
the  Hardy  Binocular  Loop,  may  be  obtained  from  F.  A. 
Hardy  &  Co.,  of  Chicago,  Illinois.  It  is  worn  like  a  pair  of  PIG.  62 

spectacles.    A  more  elaborate  binocular  magnifier,  to  be  worn 
with  an  elastic  headband,  may  be  secured  from  the  Bausch   &  Lomb 
Optical  Co.,  of  Rochester,  New  York. 

Binocular  Compound  Microscopes  with  but  a  single  objective  are  also 
obtainable.  By  means  of  a  prism  which  extends  partly  across  the  field 
about  half  of  the  light  is  directed  into  the  left  eye,  the  rest  passing  unob- 
structed to  the  right.  While  permitting  of  higher  magnification,  this  type 
of  binocular  is  not  as  universally  serviceable  as  the  other  form  described. 
The  image  is  not  erected. 

Brownian  Movement  or  Pedesis. — An  oscillating  or  dancing  motion 
observable  in  small  particles  in  a  liquid  when  seen  under  the  microscope. 

Calibration  of  Microscope. — See  "Micrometer"  (p.  197). 

Camera  Lucida. — An  apparatus  containing  a  glass  prism  or  thin  glass 
plate  so  arranged  that  when  it  is  placed  over  the  eyepiece  of  the  microscope, 
the  observer  may  see  the  image  of  the  object  under  the  microscope  projected 
on  to  his  drawing-paper  on  the  table.  The  point  of  the  pencil  is  also  visible; 
consequently  the  outline  of  the  object  may  be  readily  traced  on  the  paper. 
In  the  simpler  camera  lucidas  a  thin  neutral-tint  glass  slip  is  so  arranged 
that  it  is  in  alignment  with  the  eye-lens  of  the  ocular,  except  that  it  sets  at 


190 


Animal  Micrology 


an  angle  of  45  degrees  to  it.  When  the  microscope  is  tilted  into  a  horizontal 
position  the  observer  sees  the  image  of  the  object  reflected  from  the  upper 
side  of  the  glass  slip,  but,  since  the  latter  is  somewhat  transparent,  he  also 
sees  the  white  paper  spread  below  on  the  table  (Fig.  63). 

Another  form  of  simple  camera  lucida  is  the  Wollaston.  To  use  it  the 
microscope  must  be  inclined.  The  essential  part  of  the  camera  consists 
of  a  quadrangular  prism.  The  eye  of  the  observer  is  so 
placed  over  the  edge  of  the  prism  as  to  receive  rays  of 
light  from  the  object  with  one  portion  of  the  pupil,  and 
from  the  drawing-paper  with  the  remainder. 

Some  form  of  the  Abbe  camera  lucida,  however,  is 
used  by  most  workers.  It  consists  of  a  cap  which  is  fitted 
immediately  above  the  eyepiece  and  which  contains  two 
right-angle  prisms  cemented  together  to  form  a  cube 
(Fig.  64).  The  lower  one  of  the  prisms  is  silvered  along  its  cemented  surface, 
although  a  small  central  opening  is  left  through  which  the  object  under  the 
microscope  may  be  viewed;  connected  with  the  cap  is  an  arm  which  bears 
a  mirror,  and  this  mirror  may  be  so  adjusted  as  to  reflect  the  image  of  the 
drawing-paper  on  the  table  on  to  the  prisms  from  one  side.  The  prisms 


FIG.  63. — Simple 
Camera  Lucida. 


PIG.  64. — Abbe  Camera  Lucida 


are  so  set  that  the  silvered  surface  of  the  lower  one  reflects  this  image  upward 
to  the  eye  of  the  observer  which  also,  coincidentally,  is  viewing  the  magnified 
image  of  the  object  through  the  hole  in  the  silvering.  When  proper  adjust- 
ment of  the  light  received  from  object  and  paper  respectively  is  made,  a 
pencil-point  may  be  distinctly  seen  when  brought  into  the  field  of  vision  over 
the  paper;  consequently  the  outline  of  the  object  may  be  accurately  traced. 


The  Microscope  and  Its  Optical  Principles  191 

The  secret  of  success  in  working  with  a  camera  lucida  is  to  have  the  illumina- 
tion in  the  two  fields  properly  balanced.  Small  screens  of  tinted  glass  are 
provided  with  the  instrument  for  such  regulation.  With  the  Abbe  camera 
lucida  the  microscope  may  be  used  hi  a  vertical  or  in  an  inclined  position. 
If  the  microscope  stand  is  inclined,  the  drawing-board  upon  which  the  paper 
rests  must  have  the  same  inclination,  or  the  outline  when  drawn  will  be  dis- 
torted. Likewise,  if  the  mirror  of  the  camera  is  at  any  other  angle  than 
45  degrees,  an  adjustment  of  the  drawing-surface  must  be  made;  in  short, 
the  axial  ray  of  the  image  and  the  drawing-surface  must  always  be  at  right 
angles  to  prevent  distortion.  This  means  that  if  the  mirror  is  depressed 
below  45  degrees  the  drawing-surface  must  be  tilted  toward  the  microscope 
twice  as  much  as  the  mirror  is  depressed.  For  example,  if  the  mirror  is 
depressed  to  37  degrees  (8  below  45  degrees),  the  drawing-board  must  be 
tilted  (raised)  16  degrees.  See  also  remarks  under  "Micrometer"  (p.  197). 
When  the  camera  is  in  proper  position  the  field  of  the  microscope  should 
appear  at  about  the  same  size  as  without  the  camera.  If  the  field  is  reduced 
or  unevenly  lighted,  the  camera  is  too  near  or  too  far  from  the  ocular,  or  it 
is  tilted,  or  the  prism  is  not  properly  centered. 

Compensating  Ocular. — A  specially  designed  eyepiece  for  use  with 
apochromatic  lenses.  It  was  found  advantageous  to  undercorrect  the 
objective  and  then  to  rectify  the  aberration  by  overcorrecting  the  ocular. 
The  so-called  searching  ocular  is  a  low-power  compensating  ocular  used  for 
the  first  finding  of  objects.  The  object  once  located  in  the  field,  the  higher 
working  oculars  are  used  in  observation. 

Condenser. — A  lens  or  a  series  of  lenses  mounted  in  a  substage  attach- 
ment for  the  purpose  of  concentrating  light  upon  the  object  to  be  examined. 
They  are  made  in  various  grades  of  excellence,  non-achromatic,  achromatic, 
and  apochromatic.  Some  wide-angle  condensers  are  used  as  immersion 
condensers;  the  immersion  fluid  is  placed  between  the  upper  surface  of  the 
condenser  and  the  lower  surface  of  the  object-slide.  Condensers  are  espe- 
cially valuable  with  high-power  objectives  and  oil-immersion  lenses.  For 
the  best  results  the  condenser  must  be  accurately  centered  and  the  object 
must  lie  at  the  apex  of  the  cone  of  light  formed  by  it.  Unintelligent  use  of 
the  condenser  is  a  very  common  fault.  Condensers  are  constructed  to 
receive  parallel  rays  of  light,  hence  the  plane  mirror  only  should  be  used  with 
them  if  the  illumination  is  from  daylight.  See  "Illumination"  (p.  194). 

Correction  Collar. — A  device  for  adjusting  the  distance  between  the 
lens  systems  of  objectives  so  that  the  proper  corrections  may  be  made  for 
different  thicknesses  of  cover-glass.  Low-power  objectives  are  not  so 
sensitive  as  those  of  high  power  to  the  influence  of  the  cover-glass.  Ordinary 
objectives,  however,  are  mounted  in  a  rigid  setting  and  corrected  for  a  specific 
tube-length  and  a  standard  cover-glass  (about  0. 18  mm.  thick;  i.e.,  a  No.  2). 
With  a  cover-glass  of  different  thickness  correction  should  be  made  by 


192  Animal  Micrology 

altering  the  tube-length  of  the  microscope,  lengthening  it  for  a  thinner  cover 
and  shortening  it  for  a  thicker  one.  With  a  4  mm.  (ir  in.)  or  a  3  mm.  (i  in.) 
dry  objective  a  deviation  of  as  little  as  0 . 05  mm.  in  the  thickness  of  the  cover- 
glass,  if  uncorrected,  is  sufficient  to  obliterate  fine  details  of  the  object.  With 
homogeneous  immersion  lenses  the  defect  caused  by  different  thicknesses 
of  cover-glass  disappears  (see  "Immersion  Objective,"  p.  195).  See  also 
'Tube-Length"  (p.  201). 

Cover-Glass  Correction,  Cover-Glass  Thickness. — See  "Correction 
Collar"  (p.  191). 

Daylite  Glass. — A  specially  constructed  glass  which,  when  used  as  a 
screen  with  a  nitrogen-filled  tungsten  lamp,  yields  a  light  almost  exactly 
like  daylight.  It  gives  very  nearly  true  color  values.  The  light  is  soft 
like  that  from  a  white  cloud,  and  is  more  comfortable  to  work  with  in  micros- 
copy than  any  other  form  of  artificial 
illumination.  See  Gage,  Science, 
XLII  (October,  1915),  534,  for  a 
fuller  description. 

Dark-Ground  lUumination. — See 
"Ultramicroscopy"  (p.  201). 

Demonstration  Microscope. — A 
microscope  designed  to  be  passed 
around  a  class  with  specimen  in 
place.  Most  of  the  compound  types 
are  in  the  form  of  adjustable  tubes 

which>    when   in   use>   are   Pointed 
toward  a  window  or  a  lamp. 

Demonstration  or  Pointer  Ocular. — An  ocular  provided,  at  the  point 
where  the  real  image  of  the  object  is  produced,  with  a  delicate  rod  of  some 
kind  which  may  be  rotated  to  point  out  objects  in  the  field.  A  simple  type 
may  be  made  by  cementing  a  hair  across  the  opening  of  the  ocular  diaphragm 
with  balsam.  When  the  balsam  is  hard  the  hair  is  cut  at  the  center  of  the 
opening  and  one  end  is  removed.  It  is  necessary  to  have  both  ends  of  the 
hair  supported  until  the  balsam  hardens,  otherwise  the  free  ends  will  sag 
and  not  be  in  focus.  To  use,  rotate  the  ocular. 

A  double  demonstration  eyepiece,  by  means  of  which  the  image  formed 
by  the  objective  can  be  viewed  simultaneously  by  two  observers,  has  also 
been  devised. 

Diaphragm. — Opaque  plates  with  openings  of  various  sizes  for  regulating 
the  illumination  of  the  object  to  be  examined.  The  iris  diaphragm  (Fig.  66) 
is  the  best  type.  It  consists  of  a  series  of  overlapping  plates  placed  around 
a  central  opening  the  size  of  which  may  be  varied  by  means  of  a  lever. 
Revolving  diaphragms  are  commonly  used  on  the  cheap  grades  of  micro- 
scopes. They  consist  of  round  disks  perforated  by  openings  of  various  sizes 


The  Microscope  and  Its  Optical  Principles 


193 


which  may  be  rotated  between  the  mirror  and  the  object.  The  nearer  to  the 
object  the  diaphragm  is  placed  the  better  the  intensity  of  the  illumination 
can  be  regulated.  Most  of  the  better  class  of  microscopes  are  provided  with 
two  iris  diaphragms,  one  beneath  the  condenser  to  be  employed  when  the 
latter  is  in  use,  the  other  flush  with  the  stage  to  be  used  only  when  the  con- 
denser is  out.  If  this  second  iris  diaphragm  is  lacking,  its  place  is  taken  by 
means  of  a  cap-diaphragm  which  may  be  fitted  into  the  substage  in  the  place 
of  the  condenser.  A  central-stop  diaphragm  is  one  with  an  opaque  center 
and  a  slit  around  the  edge,  so  arranged  that  a  hollow  cone  of  light,  consisting 
of  rays  of  great  obliquity,  will  be  produced. 


FIG.  66. — Top  View  of  a  Substage  Attachment  with  Condenser  and  Lower  Iris 
Diaphragm  Thrown  out  of  Optical  Axis. 

Dissecting  Microscope. — An  instrument  so  constructed  as  to  enable  an 
operator  to  carry  on  minute  dissections  under  magnification.  Ordinarily 
they  are  simple  microscopes  mounted  on  a  stand  of  some  kind.  The  best 
instruments  (Fig.  67)  are  provided  with  well-corrected  lenses,  with  glass 
stage,  mirror,  black-and-white  substage  plate,  and  rests  for  the  hands. 
See  also  Figs.  68  and  69  for  modified  forms.  The  binocular  type,  Fig.  61,  is 
indispensable  for  the  finer  modern  technique.  See  "Binocular  Microscope." 

Embryograph. — A  form  of  camera  lucida  for  drawing  at  slight  magni- 
fication small  objects,  such  as  embryos.  A  camera  lucida  attached  to  a 
simple  microscope  is  frequently  used  for  this  purpose. 

Eye-Point. — The  point  above  an  ocular  or  lens  at  which  the  largest 
number  of  rays  from  the  instrument  enter  the  eye.  The  largest  field  of 
the  microscope  is  visible  from  this  point. 

Flatness  of  Field.— See  "Aplanatism"  (p.  187). 

Homogeneous  Immersion  Objective. — See  "Immersion  Objective" 
(p.  195). 


194 


Animal  Micrology 


Huygenian  Ocular. — See  p.  180. 

Illumination. — Any  means  employed  to  direct  light  upon  the  object 
under  observation.  Light  which  traverses  the  object  is  said  to  be  transmitted 
light.  Most  microscopical  work  in  biology  is  done  by  means  of  transmitted 
light,  hence  the  object  must  be  rendered  more  or  less  transparent  if  not 
naturally  so.  If  the  object  is  symmetrically  lighted,  the  lighting  is  desig- 
nated as  axial  or  central  illumination.  If  one  side  is  lighted  more  than 
another,  the  term  oblique  illumination  is  employed.  In  the  case  of  trans- 
mitted light,  the  light  which  traverses  the  object  is  usually  light  reflected 
from  a  mirror  because  it  is  generally  inconvenient  or  impossible  to  hold  the 
instrument  directly  toward  the  source  of  light.  Makers  of  microscopical 


PIG.  67. — Dissecting  Microscope 

appliances,  however,  now  supply  admirable  miniature  electric  lamps  which 
may  be  used  with  the  mirror  or  in  place  of  it  (Fig.  65). 

Light  which  falls  upon  the  object  and  is  reflected  from  it  to  the  eye, 
either  directly  or  through  a  microscope,  is  termed  reflected  light.  Such 
illumination  is  employed  but  little  in  ordinary  histological  work,  but  it  is 
useful  in  the  examination  of  opaque  objects  such  as  metals,  insects,  etc. 
The  illumination  may  be  increased  by  means  of  a  bull's-eye  condenser  or  a 
mirror.  In  some  microscopes  the  mirror  can  be  swung  above  the  stage  for 
the  purpose  of  illumining  an  object  which  is  to  be  studied  by  reflected  light. 


The  Microscope  and  Its  Optical  Principles  195 

The  best  light  for  microscopical  work  is  light  reflected  from  white  clouds. 
Direct  sunlight  is  never  used.  The  light  should  come  from  in  front  of  the 
observer  or  from  one  side.  Various  kinds  of  artificial  light  are  used  for  micro- 
scopical work,  such  as  an  ordinary  lamp  with  flat  wick,  the  Welsbach,  or 
the  ordinary  electric  light.  Some  of  the  newer  electric  lights  especially 
designed  for  microscopy  are  excellent.  Whatever  the  source,  the  rays 
must  be  steady  and  brilliant.  If  a  lamp  with  flat  wick  is  used,  greater  bril- 
liancy is  secured  when  the  edge  of  the  flame  is  turned  toward  the  microscope; 
the  object  should  be  lighted  directly  by  the  image  of  the  flame.  To  do  this 
with  low  powers,  the  lamp  may  have  to  be  turned  so  that  the  flame  is  oblique 
to  the  microscope. 

In  artificial  light  the  rays  are  divergent,  not  parallel,  as  in  the  case  of 
sunlight;  hence  they  will  not  come  to  focus  at  the  same  point  when  reflected 
from  the  mirror  as  the  latter  do.  This  should  be  corrected  by  using  a  large 
bull's-eye  condenser  between  the  source  of  light  and  the  mirror,  or  by  sliding 


PIG.  68. — Lens  Holder  with  Flexible  Arm 

the  mirror  along  the  mirror-bar  farther  away  from  the  stage  so  that  the  con- 
cave mirror  will  have  a  longer  distance  in  which  to  bring  the  rays  to  focus.  If 
a  substage  condenser  is  used,  the  same  results  may  be  obtained  by  depressing 
the  condenser  somewhat  below  the  level  of  the  stage.  Lamps  made  for  the 
microscope  often  have  a  metal  chimney  with  a  bull's-eye  in  one  side. 

The  objectionable  yellowness  of  most  artificial  light  may  be  eliminated 
by  interposing  a  piece  of  green  signal-glass  between  the  lamp  and  the  micro- 
scope. With  most  microscopes,  round  slips  of  blue  glass  which  fit  into  the 
substage  mechanism  are  supplied  for  this  purpose.  Many  workers  still 
employ  as  a  screen  an  ammonia  sulphate  of  copper  solution  in  a  globular 
flask.  To  make  the  solution,  dissolve  a  small  amount  of  copper  sulphate  in 
water  and  add  ammonia.  At  first  a  precipitate  appears,  but  if  an  excess 
of  ammonia  is  added  this  is  dissolved  and  a  transparent  deep-blue  liquid 
results.  This  should  be  diluted  with  water  sufficiently  to  get  a  blue  of  just 
the  proper  depth  to  render  the  transmitted  light  white  as  seen  through  the 
microscope.  The  globular  flask  also  acts  as  a  condenser. 

Immersion  Objective. — A  kind  of  objective  in  which  a  liquid  is  used 
between  the  front  lens  and  the  cover-glass.  Cedar  oil  is  the  most  widely 


196  Animal  Micrology 

used  medium.  Inasmuch  as  the  optical  properties  of  cedar  oil  (refraction 
and  dispersion)  are  almost  the  same  as  crown  glass,  it  is  often  termed  a 
homogeneous  immersion  fluid.  A  homogeneous  immersion  lens,  therefore, 
would  be  one  intended  for  use  with  such  a  fluid.  The  advantage  of  an 
immersion  over  a  dry  lens  lies  in  the  fact  that,  other  things  being  equal,  after 
leaving  the  cover-glass  rays  which  would  be  so  refracted  in  a  rarer  medium 


PIG.  69. — High-Power  Dissecting  Lens 

This  lens,  represented  as  in  use  on  the  stand  of  a  dissecting  microscope,  is  provided 
with  two  double  reflecting  Porro  prisms  which  erect  the  image  so  that  the  operator  works 
as  with  a  simple  lens. 

like  air  as  to  miss  the  front  lens  of  the  objective  reach  this  lens  in  the  case  of 
immersions  and  traverse  the  objective.  With  homogeneous  immersions  the 
rays  of  light  are  carried  without  deflection  through  cover-glass  and  fluid 
and  into  the  glass  of  the  front  lens.  Water  has  a  greater  density  than  air 
and  less  than  glass;  hence,  with  a  water  immersion  more  rays  of  light  reach 
the  front  lens  than  with  a  dry  lens,  and  less  than  with  a  homogeneous  immer- 


The  Microscope  and  Its  Optical  Principles 


197 


FIG.  70. — Diagram  to  Illus- 
trate  the  Relative  Amounts  of 
Light  Utilized  with  Dry,  Water 
Immersion,  and  Homogeneous 
Oil  Immersion  Objectives  Re. 
spectively  (after  Bausch). 


sion  lens  (Fig.  70).  The  effect  of  an  immer- 
sion is  practically  to  widen  the  angle  of  the 
lens  (see  "Angular  Aperture,"  p.  187).  The 
value  of  the  immersion  objective  is  enhanced 
if  the  immersion  fluid  is  placed  between  the 
upper  lens  of  the  condenser  and  the  slide 
as  well  as  between  the  objective  and  the 
cover-slip. 

Magnifying  Power. — The  power  of  a  lens 
to  multiply  the  apparent  dimensions  of  an 
object  viewed  through  it.  It  should  be 
expressed  in  diameters,  not  in  areas.  While 
magnifying  power  is  very  important,  it  is 
only  so  in  connection  with  resolving  power. 
If  high  power  were  the  only  essential,  a  series 
of  single  lenses  might  be  used.  The  impossi- 
bility of  using  such  a  series  for  high  magnifi- 
cation is  due  to  the  fact  that  proper  correction 
of  aberrations  cannot  be  made,  and,  conse- 
quently, a  distinct  image  cannot  be  obtained. 
For  determination  of  magnification  see 
"Micrometer"  (p.  197). 

Mechanical  Stage. — A  stage  attachment  (Fig.  71)  for  the  more  accurate 
manipulation  of  an  object  or  a  series  of  objects  which  must  be  moved  about 

under  the  objective.  The  best 
mechanical  stages  are  provided 
with  scales  and  verniers  so  that 
an  object  once  recorded  may  be 
easily  found  again.  They  are 
often  very  serviceable,  especially 
with  high  powers. 

Micrometer. — A  scale  for 
measuring  objects  under  the 
microscope.  The  stage  micrometer 
consists  of  a  finely  divided  scale 
(-rV  and  yitf  mm.)  ruled  on  glass 
or  metal.  It  is  commonly 
mounted  on  a  glass  slide  of 
standard  size.  To  determine  the 
actual  size  of  an  object  with  the 
stage  micrometer,  it  is  most  con- 
venient to  use  a  camera  lucida. 
FIG.  71. — Attachable  Mechanical  Stage  —,,  ,,.  .  ,,  ,  . 

for  Microscope.  Ihe  outline  of  the  object  to  be 


198  Animal  Micrology 

measured  is  projected  on  to  a  sheet  of  drawing-paper  and  marked  off. 
The  object  is  then  replaced  under  the  microscope  by  the  micrometer  and  the 
micrometer  scale  is  projected  on  to  the  paper.  Knowing  the  actual  distance 
between  the  lines  on  the  micrometer  scale,  the  magnification  as  well  as  the 
real  size  of  the  object  is  readily  calculated. 

The  size  of  the  image  projected  by  a  camera  lucida  on  to  a  piece  of 
drawing-paper  at  the  level  of  the  table,  however,  does  not  represent  the  true 
magnifying  power  of  the  microscope.  The  latter  is  really  considerably 
smaller  if  the  microscope  is  in  a  vertical  position  because  the  magnification 
of  a  lens  or  a  system  of  lenses  is  calculated  in  terms  of  the  conventional 
distance  of  vision  (250  mm.,  see  p.  183)  while  the  distance  from  the  ocular 
to  the  table  is  considerably  more  than  250mm.  Since  the  rays  of  light 
diverge  after  leaving  the  ocular,  manifestly  the  projected  image  will  be  larger 
(possibly  by  as  much  as  60  per  cent)  at  the  level  of  the  table  than  at  a  level 
just  250  mm.  from  the  point  of  emergence  of  the  rays  from  the  ocular.  To 
determine  the  actual  magnification  of  the  microscope,  therefore,  one  would 
have  to  bring  the  drawing-surface  to  within  250  mm.  of  this  point  of  emer- 
gence, sketch  the  projected  scale  of  the  stage  micrometer  on  the  paper,  and 
then,  by  means  of  an  ordinary  metric  rule,  compute  the  number  of  times  the 
divisions  of  the  micrometer  scale  have  been  magnified.  The  standard 
distance  of  250  mm.,  if  the  Abbe  camera  lucida  is  used  (with  camera  mirror 
at  45  degrees),  includes  the  distance  along  the  mirror-bar  from  the  optical 
axis  of  the  ocular  to  the  mirror,  plus  the  distance  from  the  mirror  to  the 
drawing-surface . 

In  practical  work  it  is  not  necessary  to  make  drawings  or  measure- 
ments exactly  at  this  standard  distance;  one  needs  only  to  have  a  scale 
made  out  for  the  distance  from  the  camera  lucida  at  which  the  drawings  are 
actually  to  be  made,  although  it  must  be  carefully  borne  in  mind  that  any 
variation  in  the  elevation  of  the  drawing-surface  will  alter  the  size  of  the 
projected  image.  A  series  of  carefully  prepared  scales  for  various  combina- 
tions of  objectives  and  oculars  should  be  made  and  kept  for  future  use. 
On  each  should  be  recorded  the  tube-length  used,  the  number  of  the  objec- 
tive and  of  the  ocular,  the  length  of  the  camera  mirror-bar,  and  the  angle 
of  the  mirror,  for  if  any  one  of  these  is  changed  the  scale  is  no  longer 
accurate. 

When  much  measuring  is  to  be  done  an  ocular  micrometer  is  used.  It 
consists  of  a  circular  glass  disk  with  a  scale  ruled  on  it  and  is  inserted  in  the 
ocular  between  the  eye-lens  and  the  field-lens.  By  means  of  a  stage  microm- 
eter the  value  of  the  divisions  of  the  ocular  micrometer  is  determined  for  a 
known  tube-length  and  every  combination  of  lenses  it  is  desired  to  use  in 
the  work  of  measurement.  Suppose  that  it  takeg  four  divisions  of  the  ocular 
micrometer  to  correspond  to  one  of  the  finer  divisions  of  the  stage  micrometer, 
then  since  the  divisions  of  the  latter  are  equal  to  TOT  mm.,  each  space  in  the 


The  Microscope  and  Its  Optical  Principles 


199 


ocular  micrometer  must  be  equal  to  TOT  mm.,  that  is,  0.0025  mm.  A  filar 
or  screw  micrometer  is  a  more  convenient  form  of  ocular  micrometer,  which  is 
provided  with  delicate  movable  spider  lines  that  can  be  adjusted  to  the  space 
to  be  measured  by  means  of  a  fine  screw  with  very  accurately  cut  threads 
(Fig.  72).  At  the  end  of  the  screw  is  a  graduated  disk  which  gives  the  value 
of  the  distance  between  the  spider  lines.  The  pitch  of  the  screw  is  either 
aV  inch  or  0 . 5  mm.  When  once  the  valuation  of  this  ocular  micrometer  has 
been  determined  by  means  of  a  stage  micrometer,  measurements  can  be 
made  rapidly  and  with  great  precision. 

The  step  micrometer,  in  which  the  intervals  are  arranged  in  groups  of 
ten,  each  group  being  conspicuously  marked  by  a  black,  stairlike  notching 
along  one  side,  is  one  of  the  most  desirable  types  of  ocular  micrometers. 

Micron. — The  one-thousandth  pa'rt  of  a  millimeter;    expressed  briefly 
by  the  Greek  letter  /x. 
It  is  the  unit  of  meas- 
urement in  microscopy. 

Mirror. — The  com- 
pound microscope  is 
usually  provided  with 
both  concave  and  plane 
mirrors,  which  may  be 
rotated  or  swung  in  any 
direction.  The  plane 
mirror  is  used  with  the 
condenser;  the  concave, 
whenever  it  is  of  advan- 
tage to  have  light  con- 
centrated  upon  the 
object,  with  the  con- 
denser out.  The  mirror  should  be  capable  of  being  moved  up  or  down  the 
mirror-bar  so  that  it  can  be  accurately  focused  upon  the  object.  See  also 
"Illumination"  (p.  194). 

Muscae  Volitantes. — Small  filaments  or  specks  which  float  across  the 
field  of  vision.  They  are  really  small  opacities  in  the  vitreous  humor  of 
the  eye. 

Numerical  Aperture. — A  system  which  expresses  the  efficiency  of  an 
objective  by  indicating  the  relative  proportion  of  light  rays  which  traverse 
it  to  form  an  image.  With  the  introduction  of  immersion  objectives,  it 
became  evident  that  angular  aperture  alone  is  not  sufficient  to  indicate  the 
real  capacity  of  an  objective.  For  instance,  an  immersion  and  a  dry  lens 
may  be  of  precisely  the  same  angular  aperture,  and  yet  the  immersion  lens 
is  more  efficient  because  it  sends  more  rays  of  fight  through  the  objective 
(see  "Immersion  Lens,"  p.  195).  It  was  found  necessary  to  take  cognizance 


FIG.  72. — Filar  Micrometer 


200  Animal  Micrology 

of  the  medium  which  intervenes  between  the  cover-glass  and  the  front  lens 
of  the  objective. 

Professor  Abbe,  in  1873,  proposed  the  name  numerical  aperture  and 
introduced  the  formula  N.A.  =  n  sin  u,  in  which  n  signifies  the  refractive 
index  of  the  medium  between  cover-glass  and  objective,  and  u  equals  half 
the  angle  of  aperture.  That  is,  by  multiplying  the  refractive  index  of  the 
medium  by  the  sine  of  half  the  angle  of  aperture  the  numerical  aperture  is 
obtained.  For  example,  suppose  that  one  had  an  oil-immersion  lens  of 
90  degrees  angular  aperture,  then  half  the  angle  of  aperture  is  45  degrees, 
and  by  turning  to  a  table  of  natural  sines,  the  sine  of  45  degrees  is  found 
to  be  0 . 707.  The  refractive  index  of  cedar  oil  is  1 . 52.  Then  N.A.  =  1 . 52X 
0.707=1.075.  Suppose  that  the  lens  were  a  dry  instead  of  an  immersion 
lens;  then  since  the  refractive  index  of  air  is  1,  the  formula  would  read 
N.A.=  1X0. 707=0.707.  Thus  the  two  products  1.075  and  0.707,  respec- 
tively, represent  the  relative  capacities  of  an  oil  immersion  and  a  dry  objec- 
tive of  90  degrees  angular  aperture. 

Parfocal. — A  term  ordinarily  applied  to  eyepieces  of  different  powers 
that  may  be  exchanged  in  the  microscope  without  very  materially  affecting 
the  focus  of  the  instrument.  The  term  is  also  applied  to  objectives  attached 
to  a  revolving  nosepiece  if  each  is  approximately  in  focus  when  turned  into 
place. 

Pedesis. — Same  as  Brownian  movement. 

Penetration. — The  quality  of  an  objective  that  permits  of  "looking 
into"  an  object  having  sensible  thickness.  It  is  greatest  with  low  powers 
and  narrow  angles  and  is  antagonistic  to  resolving  power.  It  is  the  natural 
consequence  of  certain  conditions  in  the  making  of  lenses  and  is  reckoned 
of  secondary  importance,  because  practically  the  same  results  are  obtained 
by  manipulating  the  fine  adjustment. 

Photomicrography. — Photography  of  small  or  microscopic  objects. 
The  subject,  although  of  great  importance,  is  too  extensive  to  enter  into  in 
the  brief  space  that  could  be  allotted  to  it  in  an  elementary  treatise  such  as 
this.  (See,  however,  p.  150.)  An  excellent  chapter  on  photomicrography 
and  a  bibliography  will  be  found  in  Gage's  The  Microscope. 

Pointer  Ocular.— See  "Demonstration  Ocular"  (p.  192). 

Polariscope. — As  used  in  microscopy  the  polariscope  consists  of  two 
parts,  each  composed  of  a  Nicol  prism  of  Iceland  spar;  one,  the  polarizer, 
fits  into  the  substage,  and  the  other,  the  analyzer,  is  inserted  between  the 
objective  and  the  tube  of  the  microscope  or,  in  some  forms,  just  above  the 
ocular.  The  polariscope  is  used  more  in  chemical  and  in  geological  than  in 
histological  work.  Some  of  the  uses  are  as  follows:  determining  whether 
an  object  is  singly  or  doubly  refractive;  detecting  the  presence  of  minute 
crystals;  determining  the  composition  of  rocks;  'examining  sections  of  bone, 


The  Microscope  and  Its  Optical  Principles  201 

hoof  and  horn,  hairs  and  fibers  of  animals  and  plants,  starch,  etc.,  for  certain 
characteristic  and  striking  effects. 

Projection  Ocular. — An  ocular  specially  designed  for  projecting  a  micro- 
scopic object  on  to  a  screen  or  for  use  in  microphotography.  While  ordi- 
narily used  with  apochromatic  objectives,  they  may  be  used  with  ordinary 
objectives  of  large  numerical  aperture.  The  eye-lens  is  movable  so  that  a 
sharp  focus  (indicated  by  a  distinct  image  of  the  diaphragm)  may  be  obtained 
at  different  screen  distances. 

Resolving  Power. — The  quality  of  an  objective  which  enables  the 
observer  to  make  out  fine  details  of  structure.  It  is  the  most  essential 
property  for  precision  in  observation,  and  determines  largely  the  excellence 
of  an  objective.  Resolving  power  depends  upon  careful  correction  of 
aberrations,  general  accuracy  in  the  mechanical  construction  of  the  micro- 
scope, and  upon  the  aperture  of  the  objective  (see  "Angular  Aperture," 
"Numerical  Aperture,"  pp.  187,  199).  Resolving  power  is  tested  by  the 
resolution  of  fine  parallel  lines  ruled  on  glass  or  the  striae  on  the  surface  of 
diatoms.  The  test  is  to  determine  how  many  lines  to  the  inch  or  centimeter 
may  be  distinguished,  and  whether  the  objective  simply  glimpses  the  r^ark- 
ings  or  whether  it  resolves  them  clearly.  The  wider  the  angle  of  aperture 
the  better  the  resolving  power,  provided  the  width  is  not  so  great  as  to  inter- 
fere with  the  correction  of  the  lenses.  The  increased  resolution  of  immer- 
sion lenses  is  due  to  the  fact  that  the  immersion  fluid  practically  widens 
the  angle  of  aperture  (see  "Immersion  Objective,"  p.  195). 

Tube-Length. — The  distance  between  the  places  of  insertion  of  ocular 
and  objective  into  the  tube  of  the  microscope.  There  are  two  standard 
tube-lengths;  the  short  standard  is  160  mm.  (6-ft-  inches),  the  long  standard, 
216  mm.  (8?V  inches).  Some  makers,  however,  do  not  adhere  to  the  stand- 
ards. The  optical  efficiency  of  the  instrument  is  the  same  in  either  case. 
The  short  length  is  more  advantageous  in  that  it  is  more  compact.  The 
lenses  must  be  corrected  for  the  length  of  tube  with  which  they  are  to  be 
used.  The  short  standard  is  in  use  in  most  American  laboratories. 

Ultramicroscopy.. — A  system  of  microscopical  inspection  in  which  objects 
are  examined  by  reflected  light.  The  object  appears  to  be  self-luminous 
against  a  dark  field,  hence  the  term  dark-ground  illumination  is  often  used  as 
descriptive  of  the  method.  Objects  to  be  studied  in  this  way  are  usually 
semi-transparent  or  consist  of  fine  particles  such  as  occur  in  colloidal  suspen- 
sions. In  lighting,  rays  of  great  obliquity  are  used  so  that  only  such  traverse 
the  objective  as  are  deflected  from  some  object  in  the  field.  The  great  value 
of  the  method  lies  in  the  fact  that  particles  may  be  rendered  visible  which  are 
wholly  invisible  with  the  microscope  as  ordinarily  used. 

For  low  powers  without  a  condenser,  the  diaphragm  must  be  wide  open 
and  the  mirror  so  tilted  that  the  object  is  lighted  by  oblique  rays  which 


202  Animal  Micrology 

cannot  get  directly  into  the  front  lens  of  the  objective.  With  a  condenser, 
a  central-stop  diaphragm  is  used  which  admits  only  marginal  rays.  By 
making  an  ordinary  diaphragm  eccentric,  somewhat  the  same  effect  may  be 
secured.  For  practice,  place  a  drop  of  10  per  cent  solution  of  salicylic 
acid  in  95  per  cent  alcohol  on  a  slide  and  leave  it  until  the  alcohol  evaporates. 
Examine  the  residue  of  crystals  in  the  ordinary  way  and  then  by  dark- 
ground  illumination.  By  the  latter  method  the  crystals  should  appear  bril- 
liantly lighted  on  a  dark  background.  Add  a  small  drop  of  the  solution  to  the 
crystals  and  watch  crystallization  under  dark-ground  illumination. 

For  high  powers,  according  to  one  system,  very  wide  apertures  (greater 
than  1.00  N.A.;  see  p.  199)  are  necessary  in  the  condensers.  Some  makers 
(e.g.,  Leitz,  Reichert)  use  specially  modified  condensers.  Others  (e.g.,  Beck, 
Siedentopf)  substitute  a  parabolic  reflector  for  the  condenser.  In  the 
method  of  Siedentopf  and  Zsigmondy,  the  field  is  lighted  from  one  side,  at 
right  angles  to  the  axis  of  the  microscope,  by  a  wedge  or  cone  of  bright 
light.  In  another  method,  useful  for  both  high  and  low  powers,  an  objective 
of  wide  aperture  and  a  condenser  of  moderate  aperture  are  employed.  The 
field 's  lighted,  as  in  ordinary  microscopy,  by  a  cone  of  light  from  the  conden- 
ser, but  a  diaphragm  or  stop  of  the  right  size  to  cut  out  the  central  rays  of  light 
is  placed  on  the  back  lens  of  the  objective.  In  this  way  only  those  rays  which 
have  entered  the  marginal  zones  of  the  objective  pass  on  to  form  an  image, 
and  among  these  are  the  rays  which  have  been  deflected  by  objects  in  the 
field.  A  somewhat  similar  result  may  be  attained  by  using  a  stop  in  the 
eye-point  (p.  193).  For  a  fuller  discussion  of  ultramicroscopy  and  dark- 
ground  illumination  see  A.  E.  Wright,  Principles  of  Microscopy,  chap.  xiv. 

Overcorrection  and  Undercorrection. — In  correcting  for  chromatic 
aberration,  if  the  concave  lens  is  stronger  than  is  necessary  to  neutralize  the 
aberration  of  the  convex  lens,  the  blue  rays  are  brought  to  focus  beyond  the 
true  principal  focus  of  the  objective,  and  the  latter  is  said  to  be  overcorrected; 
if  the  concave  lens  is  not  strong  enough,  the  result  is  what  is  known  as  under- 
correction.  In  case  of  overcorrection,  the  object  takes  on  an  orange  tint 
if,  after  focusing,  the  distance  between  object  and  objective  is  slightly 
increased;  or  it  becomes  of  bluish  color  if  the  distance  is  decreased.  In 
case  of  undercorrection,  just  the  reverse  is  true.  In  some  instances  the  objec- 
tive is  purposely  undercorrected,  and  the  eyepiece  (e.g.,  compensating  ocular) 
is  equally  overcorrected. 

Working  Distance. — The  distance  between  the  front  lens  of  the  objec- 
tive and  the  object  when  the  latter  is  in  focus.  With  high  powers  it  is  very 
small,  so  that  with  some  oil-immersion  objectives,  if  a  thick  cover  is  used, 
it  is  impossible  to  focus  upon  the  object.  For  this  reason  thin  cover-glasses 
(No.  1)  should  be  used  on  preparations  which  are  to  be  used  with  high-power 
immersion  lenses.  For  examination  under  high-power  dry  lenses,  however, 
see  remarks  under  "Correction  Collar"  (p.  191). 


The  Microscope  and  Its  Optical  Principles  203 

MANIPULATION  OF  THE  COMPOUND  MICROSCOPE 

1.  Always  handle  the  instrument  cautiously;    it  is  a  delicate 
mechanism.     Lift  it  by  the  base  or  by  a  handle  specially  provided, 
never  by  the  tube. 

2.  The  work-table  should  be  of  such  a  height  that  the  observer 
can  sit  at  it  comfortably  without  compressing  the  chest  or  tiring 
the  neck.     Sit  as  upright  as  possible.     If  the  instrument  is  inclined, 
it  should  set  farther  in  on  the  table  than  if  it  is  in  the  upright  position. 

3.  With  a  piece  of  old  linen,  a  chamois  skin,  or  a  bit  of  lens- 
paper,  carefully  clean  the  eyepiece  to  be  used  and  put  it  in  place. 
Always  use  the  low-power  eyepiece  first. 

4.  Likewise  clean  and  attach  the  objective  (low  power  first)  after 
elevating  the  tube  far  enough  above  the  stage  for  this  purpose. 
Guard  particularly  against  screwing  the  objective  in  crooked,  as  this 
will  injure  the  threads.     It  is  best  to  swing  the  objective  between  the 
first  and  second  fingers  of  one  hand  and  bring  the  screw  squarely  into 
contact  with  the  screw  of  the  tube  (or  nosepiece) ;  with  the  thumb 
and  forefinger  of  the  other  hand  it  is  then  screwed  into  place. 

5.  Bring  the  draw-tube  to  the  standard  length   (see  "Tube- 
Length,"  p.  201)  for  which  the  lenses  are  corrected.     If  a  nosepiece 
is  used,  allowance  must  be  made  for  its  height.     In  some  of  the  more 
recently  made  microscopes,  however,  the  scale  on  the  back  of  the 
draw-tube  includes  the  nosepiece.     In  pushing  in  or  drawing  out  the 
draw-tube  always  grasp  the  milled  head  of  the  coarse  adjustment 
also,  so  that  the  tube  as  a  whole  will  not  be  shifted. 

6.  Place  the  slide  which  bears  the  object  on  the  stage  with  the 
object  over  the  central  opening  of  the  latter,  and  clamp  it  in  place 
by  means  of  the  spring  clips.     While  looking  at  the  object  from  one 
side,  turn  the  mirror  until  a  flood  of  light  shines  up  through  the 
center  of  the  stage. 

7.  Lower  the  tube  until  the  objective  nearly  touches  the  cover- 
glass,  then  look  through  the  eyepiece  and  slowly  raise  the  tube 
by  means  of  the  coarse  adjustment  until  the  specimen  to  be  examined 
is  plainly  visible.     Focus  accurately  by  means  of  the  fine  adjustment. 
If  a  high-power  objective  is  being  used,  since  it  must  come  very 
near  the  cover,  the  operator  should  lower  his  head  to  the  level  of  the 


204  Animal  Micrology 

stage,  and  look  toward  the  light  between  objective  and  cover-glass 
in  order  to  prevent  actual  contact.  This  is  of  great  importance,  for 
otherwise  the  objective  or  the  object  is  liable  to  injury.  Remember 
that  in  focusing  up  the  lowest  part  of  the  object  comes  into  view  first, 
the  highest  part  last.  It  is  often  easier  to  locate  the  object  if  the 
preparation  is  moved  about  slightly  while  focusing. 

8.  The  higher  the  power  the  more  difficult  it  is  to  find  an  object 
or  a  particular  part  of  it.     For  this  reason  the  finding  is  usually  done 
by  means  of  a  low-power  objective,  or  a  low-power  ocular,  or  both, 
and  after  accurately  centering  the  object  in  the  field,  the  high  power 
is  attached.     In  case  a  revolving  nosepiece  is  used,  great  care  should 
be  used  in  turning  in  the  high  power  not  to  strike  the  slide  with  the 
objective.     This  is  very  likely  to  happen  if  the  objectives  are  not 
parfocal.     When  objectives  are  not  parfocal  they  may  usually  be 
made  so  by  putting  a  paper  or  bristol-board  collar  on  them. 

9.  After  the  object  is  in  focus  give  any  further  attention  to  the 
illumination  that  is  necessary  (see  "Illumination"  and  "Mirror," 
pp.  194,  199).     If  intensified  illumination  is  desired,  use  the  concave 
mirror,  or  use  the  substage  condenser  and  the  plane  mirror.     For 
ordinary  purposes  the  field  should  be  evenly  illuminated,  although 
oblique  light  is  frequently  useful.     Manipulate  the  diaphragm  until 
the  structure  to  be  studied  shows  with  the  greatest  distinctness.     Too 
much  light  "drowns"  the  object,  and  is  hard  on  the  eyes.     (To 
determine  the  proper  distance  at  which  the  concave  mirror  should 
stand  below  the  stage,  let  direct  sunlight  shine  upon  the  mirror,  and 
then  adjust  the  latter  so  that  the  apex  of  the  cone  of  light  comes  just 
at  the  top  of  the  stage  where  the  object  will  rest.) 

If  particles  of  dust  or  cloudiness  appear  in  the  field,  determine,  by 
moving  the  slide  and  rotating  the  ocular  one  after  the  other,  whether 
slide,  objective,  or  ocular  requires  further  cleaning.  A  camel's 
hair  brush  is  often  more  effective  than  lens-paper  or  cloth  in  removing 
bits  of  dust. 

10.  In  using  oil-immersion  objectives,  a  small  drop  of  cedar  oil 
(specially  prepared  by  the  maker  of  the  lens)  is  applied  to  the  front 
lens  by  means  of  a  small  rod  or  brush.     It  is  very  important  to  keep 
the  oil  free  from  dust,  and  to  see  that  it  does  not  contain  air  bubbles 


The  Microscope  and  Its  Optical  Principles  205 

when  applied  to  the  lens.  Carefully  lower  the  tube  until  the  oil  on 
the  objective  comes  in  contact  with  the  cover-glass.  The  operator 
should  lower  his  head  to  the  level  of  the  stage  to  observe  this  properly. 
Focus  up  as  with  a  dry  objective.  For  critical  work  immersion  oil 
should  also  be  placed  between  the  condenser  and  the  lower  side  of  the 
slide.  With  a  piece  of  lens-paper  or  a  soft  cloth  clean  the  immersion 
lens  immediately  after  you  have  finished  using  it.  Likewise  remove 
the  oil  from  the  cover-glass.  Oil  which  has  hardened  on  the  cover- 
glass  should  be  removed  with  lens-paper  wet  with  xylol. 

If  the  immersion  oil  becomes  too  dense,  as  is  likely  after  some 
months,  it  may  be  diluted  with  pure  cedar  oil. 

11.  The  range  of  the  fine  adjustment  is  limited.     Keep  it  as 
near  the  middle  point  as  possible.     If  the  tube  does  not  respond  to 
the  movement  of  the  screw,  you  have  probably  gone  beyond  the 
range  of  the  fine  adjustment. 

12.  In  working  with  the  microscope  keep  both  eyes  open.    The 
eye  which  is  not  in  use  soon  becomes  accustomed  to  ignoring  objects 
in  the  field  of  vision.    To  avoid  fatigue  it  is  well  to  use  first  one  eye 
and  then  the  other  for  observation.     The  eye  should  be  placed  at  the 
eye-point  (p.  193)  of  the  lens.    This  is  some  distance  from  the  eye- 
lens  in  low-power  eyepieces,  close  to  it  in  high-power  eyepieces. 

Wliile  observing,  with  one  hand  keep  the  fine  adjustment  moving. 
This  relieves  the  eye  of  the  strain  of  attempting  to  focus  on  different 
depths  of  the  object. 

13.  Put  the  microscope  in  its  case  when  you  have  finished  using  it, 
or  at  least  cover  it  with  a  cloth  or  cone  of  paper.     For  further  details 
regarding  the  use  or  care  of  the  microscope  consult  one  of  the  following 
books:    The  Microscope,  by  Gage;    Principles  of  Microscopy,  by 
Wright;    The  Microscope  .and  Its  Revelations  (1,200  pages),  by  Car- 
penter and  Dallinger. 

14.  Do  not  apply  alcohol  to  any  part  of  the  instrument.     The 
lenses  may  be  cleaned  ordinarily  by  breathing  upon  them  and  wiping 
them  with  a  rotary  motion  on  lens-paper  or  a  piece  of  soft  old  linen. 
In  case  a  solvent  must  be  used  for  balsam  or  oil,  benzene  is  the  one 
commonly  recommended.     It  must  be  quickly  wiped  away  so  that  it 
will  not  affect  the  setting  of  the  lens. 


206  Animal  Micrology 

15.  Read  carefully  in  the  catalogue  of  the  maker  of  your  instru- 
ment what  is  said  about  its  construction. 

16.  Determine  the  magnifications  of  your  various  combinations 
of  lenses  as  described  under  the  heading  of  "Micrometer"  (p.  197). 

The  beginner  in  microscopy  should  acquaint  himself  with  various 
common  objects  that  are  liable  to  get  into  his  preparations  in  the 
form  of  dust,  etc.,  so  that  he  may  not  mistake  them  for  essential  parts 
of  his  specimen.  Such  objects  are  hairs,  fibers  of  silk,  wool,  linen, 
cotton,  and  the  like,  and  particularly  air  bubbles.  Air  bubbles  are 
usually  circular  with  black  borders  and  bright  centers;  they  may 
show  tinges  of  color.  Examine  a  drop  of  saliva  for  examples. 


APPENDIX  B 

SOME  STANDARD  REAGENTS  AND  THEIR  USES 
I.    FIXING  AND  HARDENING  AGENTS 

1.  Acetic  Acid. — Acetic  acid  is  more  commonly  used  in  mixtures 
or  in  diluted  form  than  pure.     It  is  valuable  because  it  tends  to  pro- 
duce good  optical  differentiation  and  facilitates  penetration.     When 
employed  alone  it  causes  some  tissues  to  swell  and  disintegrate. 
Inasmuch  as  most  fixing  agents  give  the  best  results  when  they  have 
an  acid  reaction,  from  1  to  5  per  cent  of  acetic  acid  is  generally  added 
to  acidify  them  in  case  they  are  not  naturally  acid.    Any  reagent 
containing  a  very  large  proportion  of  acetic  acid  should  be  allowed 
to  act  for  only  a  short  time.    Acetic  acid  is  also  of  great  value  in 
mixtures  because  it  counteracts  the  shrinking  action  of  certain 
reagents.    Ordinary  acetic  acid  is  of  about  36  per  cent  strength; 
glacial  acetic,  of  about  99 . 5  per  cent  strength.     The  latter  is  meant 
when  mentioned  in  this  book  unless  otherwise  specified. 

A  strength  of  from  0 . 2  to  1  per  cent  is  recommended  by  Flemming 
for  work  on  cell  nuclei.  Strong  glacial  acetic  acid  is  sometimes  used 
for  highly  contractile  animals,  such  as  Coelenterata,  Mollusca,  and 
Vermes.  The  animal  is  rapidly  flooded  with  the  acid  and  remains 
immersed  until  it  is  thoroughly  penetrated  (6  to  10  minutes).  It 
is  then  washed  in  repeated  changes  of  50  or  70  per  cent  alcohol  and 
left  to  harden  in  70  to  83  per  cent  alcohol.  The  pure  acid,  if  allowed 
to  act  for  more  than  a  few  minutes,  swells  and  softens  the  tissues. 
Acetic  acid  should  not  be  used  when  connective  tissue  or  delicate 
calcareous  structures  are  to  be  preserved. 

2.  Acetic  Alcohol. — Carnoy  recommends  each  of  the  following 
formulae: 

a)  Glacial  acetic  acid 1  part 

Absolute  alcohol 3  parts 

6)  Glacial  acetic  acid 1  part 

Absolute  alcohol 6  parts 

Chloroform 3  parts 

207 


208  Animal  Micrology 

The  chloroform  is  said  to  hasten  the  action  of  the  mixture. 
Either  of  these  reagents  penetrates  well  and  acts  rapidly.  Solution  b 
is  especially  good  for  glandular  or  lymphatic  tissue.  Almost  any  stain 
will  follow  them.  Even  such  difficult  objects  as  the  eggs  of  Ascaris 
may  be  fixed  by  the  second  mixture.  The  reagent  should  be  washed 
out  in  absolute  or  at  least  in  strong  alcohol. 

Absolute  alcohol  to  which  20  per  cent  acetic  acid  has  been  added 
is  also  in  use  in  Boveri's  laboratory  for  Ascaris.  Material  is  left 
overnight  in  it. 

A  mixture  of  absolute  alcohol,  glacial  acetic  acid,  and  chloroform, 
equal  parts,  saturated  with  corrosive  sublimate  (formula  of  Carnoy 
and  Lebrun),  becomes  even  more  valuable  for  the  fixation  of  difficult 
objects.  According  to  Lee,  isolated  ova  of  Ascaris  are  fixed  in  30 
seconds,  entire  oviducts  in  10  minutes,  in  this  liquid.  It  is  good  for 
cytological  work  in  general. 

3.  Alcohol. — Alcohol  is  used  especially  for  gland  cells  and  for 
preserving  the  brain  and  spinal  cord  for  NissFs  method  of  staining 
nerve  cells.     See  "Alcohol  Fixation,"  p.  28;  also  reagents  1  and  2, 
p.  7,  and  memoranda  on  p.  13. 

Alcohol  and  Chloroform. — See  2,  6. 

Bensley's  Formol-Bichromate-Sublimate  Mixture. — See  p.  153. 

Bichloride  of  Mercury. — -See  "Corrosive  Sublimate." 

4.  Bichromate  of  Potassium. — Bichromate  of  potash  is  one  of 
the  oldest  and  best  known  fixing  reagents.     At  present  it  is  more  com- 
monly used  in  mixtures  than  alone.     It  is  widely  used  in  hardening 
nervous  tissue.     Its  fixation  of  nuclei  is  unsatisfactory  unless  it  is 
properly  corrected  through  the  addition  of  acetic  acid.     It  acts  very 
slowly,  about  three  weeks  being  necessary  to  harden  properly  a 
sheep's  eye,  and  from  three  to  six  months  for  a  good-sized  brain.    A 
weak  solution  (2  per  cent)  should  be  used  at  first,  to  be  replaced 
gradually  by  stronger  solutions  up  to  5  per  cent.     When  hardening 
is  completed  the  object  should  be  thoroughly  washed  in  running 
water  and  then  put  into  alcohol;    begin  with  low  percentages  of 
alcohol  and  gradually  increase  the  strength  up  to  70  or  80  per  cent. 
Change  the  alcohol  as  often  as  it  becomes  yellow.     After  the  object 
has  been  placed  in  alcohol,  keep  it  in  the  dark  in  order  to  prevent  a 


Some  Standard  Reagents  and  Their  Uses  209 

precipitate  forming  on  the  surface.  Either  carmine  or  hematoxylin 
may  be  used  as  a  stain  after  bichromate  of  potash.  In  case  carmine 
is  used,  the  staining  is  best  done  before  the  object  is  placed  in  alcohol. 
Tissues  which  do  not  stain  well  should  be  placed  for  3  hours  in  acid 
alcohol  and  then  washed  in  alcohol  before  staining. 

5.  Bichromate  of  Potassium  and  Acetic  Acid   (Tellyesnicky's 
fluid).— 

Bichromate  of  potassium 3  grams 

Glacial  acetic  acid 5  c.c. 

Water 100  c.c. 

It  is  best  not  to  add  the  acetic  acid  until  just  before  using.  This 
is  a  good  general  reagent.  It  is  valuable  for  embryos.  Objects 
should  remain  in  some  20  volumes  of  the  fluid  from  24  to  48  hours, 
according  to  size.  It  is  well  to  change  the  fluid  once  after  a  few 
hours.  After  fixation,  tissues  should  be  washed  thoroughly  in 
running  water  (6  to  12  hours)  and  passed  through  alcohols  of  increas- 
ing strength  beginning  with  15  per  cent. 

6.  Bichromate  of  Potassium  and  Corrosive  Sublimate  (Zenker's 
fluid). — For  formula,  see  p.  8,  reagent  7. 

Zenker's  is  a  valuable  reagent  for  both  histological  and  embryo- 
logical  material  (embryos  up  to  25  mm.) .  Several  hours  are  required 
for  fixation:  2  to  4  hours  for  a  2-day  chick;  8  to  10  hours  for  objects 
or  embryos  of  6  to  8  mm. ;  24  hours  for  embryos  of  12  to  14  mm.,  etc. 
For  washing,  running  water  is  employed  for  from  1 2  to  24  hours.  The 
object  is  then  transferred  to  gradually  increasing  strengths  of  alcohol 
up  to  70  per  cent,  leaving  it  according  to  size  from  1  to  3  hours  in  each 
alcohol.  To  remove  the  excess  of  corrosive  sublimate,  see  14, 
"Caution"  1.  Almost  any  stain  follows  this  reagent  well.  Both 
nuclear  and  cytoplasmic  structures  are  properly  fixed. 

7.  Bichromate  of  Potassium,  Corrosive  Sublimate,  and  Formalin 
(Zenker  formalin  mixtures). — 

A.  Hetty's  Fluid: 

Prepare  a  Zenker's  fluid,  but  instead  of  acetic  acid  add  formalin 
in  the  same  proportion  and  in  the  same  way.  Good  for  tissues  in 
which  it  is  desired  to  examine  the  granular  cytoplasmic  contents. 


210  Animal  Micrology 

B.  Danchakoff's  Mixture: 

Corrosive  sublimate 50  parts 

Potassium  bichromate 25  parts 

Sulphate  of  soda 10  or  12  parts 

Water 1,000  parts 

Boil  to  dissolve.  Just  before  using  add  sufficient  formalin  to 
make  the  solution  contain  5  per  cent  for  soft  tissues  or  10  per  cent 
for  dense  tissues.  Fix  for  from  2  to  4  hours,  never  more  than  6  hours. 
Keep  the  mixture  at  about  37°  C.  during  fixation.  This  fluid 'is 
useful  for  some  kinds  of  cytological  work. 

8.  Bichromate   of  Potassium  and   Cupric   Sulphate    (Erlicki's 
fluid).— 

Bichromate  of  potash 5  grams 

Sulphate  of  copper 2  grams 

Distilled  water 220  c.c. 

Pulverize  the  crystals  before  adding  the  water. 

Erlicki's  fluid  is  an  excellent  reagent  for  general  use,  and  is 
especially  valuable  for  voluminous  objects  such  as  advanced  embryos. 
Its  principal  drawback  is  the  length  of  time  required  properly  to 
harden  objects  (five  days  to  three  weeks).  The  process  may  be 
hastened  by  keeping  the  fluid  containing  the  tissue  at.the  temperature 
of  an  incubator  (39°  C.).  At  the  end  of  this  time  transfer  the 
object  to  35  per  cent  alcohol,  keeping  it  in  the  dark  for  two  hours  to 
avoid  precipitation.  The  alcohol  should  be  changed  occasionally 
during  this  time.  Repeat  the  process,  using  50  per  cent  alcohol,  and 
finally  preserve  the  material  in  70  per  cent  alcohol. 

9.  Bichromate  of  Potassium  and  Sodium  Sulphate   (Miiller's 
fluid).— 

Bichromate  of  potassium 20  to  25  grams 

Sodium  sulphate 10  grams 

Water 1,000  c.c. 

Muller's  fluid  is  an  old  and  widely  used  reagent.  It  is  especially 
valuable  for  the  nervous  system.  It  acts  very  slowly.  Specimens 
require  immersion  in  a  large  quantity  of  the  fluid  from  three  to  ten 
weeks,  according  to  size.  The  solution  should  be  changed  every  two 


Some  Standard  Reagents  and  Their  Uses  211 

days  for  the  first  ten  days,  and  later  about  once  a  week.  If  a  scum 
appears  at  any  time,  the  fluid  should  be  changed.  In  washing,  the 
tissues  are  placed  in  running  water  for  a  number  of  hours  and  are 
then  treated  with  gradually  increasing  strengths  of  alcohol  in  the 
usual  manner.  For  some  purposes,  however,  the  tissue  is  transferred 
directly  from  the  fluid  to  70  per  cent  alcohol.  In  any  event,  the 
material  should  always  be  kept  in  the  dark  to  prevent  precipitation. 

Bouin's  Picro-Formol. — See  pp.  9  and  29. 

Carney's  Acetic  Alcohol. — See  2. 

10.  Chloride  and  Acetate  of  Copper  (liquid  of  Ripart  and  Petit). — 

Camphor  water 75 . 0    grains 

Crystallized  acetic  acid 1.0    gram 

Distilled  water 75.0    c.c.   ' 

Acetate  of  copper 0 . 30  gram 

Chloride  of  copper 0.39  gram 

This  is  a  good  reagent  for  cytological  work  where  objects  are 
to  be  studied  in  as  fresh  a  condition  as  possible.  Methyl  green 
(reagent  60)  should  be  used  for  staining.  Only  aqueous  media  are 
employed  with  such  material. 

11.  Chromic  Acid. — Aqueous  solutions  of  from  0.2  to  1  per 
cent  are  used.     The  acid  is  best  kept  in  the  form  of  a  1  per  cent 
stock  solution.     Tissues  are  left  in  at  least  fifty  times  their  volume 
of  the  acid  for  from  24  hours  for  small  pieces  to  one  or  more  weeks  for 
larger  ones.     The  objects  are  then  washed  in  running  water  for 
several  hours,  after  which  they  are  treated  with  gradually  increasing 
strengths  of  alcohol.     Do  the  washing  and  dehydrating  in  the  dark. 
If  sections  of  chromic-acid  material  do  not  stain  readily,  they  should 
be  treated  for  three  hours  with  acid  alcohol,  washed  out  with  ordinary 
alcohol,  and  then  stained.     Hematoxylin  or  some  of  the  anilins  are 
the  best  stains  for  chromic  material.     Chromic  acid  hardens  much 
more  rapidly  than  bichromate  of  potash.     It  makes  tissues  extremely 
brittle. 

12.  Chrom-Acetic-Osmic  Acid  (Flemming's  solution).— 

Chromic  acid,  1  per  cent  aqueous  solution 15  parts 

Osmic  acid,  2  per  cent  aqueous  solution 4  parts 

Glacial  acetic  acid 1  part 


212  Animal  Micrology 

This  is  the  so-called  "strong"  solution  of  Flemming.  The 
mixture  should  not  be  made  until  immediately  before  using,  because 
it  deteriorates  if  allowed  to  stand  for  any  considerable  length  of  time. 
The  fluid  is  valuable  for  cytological  work,  especially  for  the  study 
of  karyokinetic  figures.  Only  small  pieces  of  tissue  should  be  used,  as 
the  reagent  penetrates  poorly.  They  should  remain  in  the  fluid  for 
from  24  to  48  hours  and  then  be  washed  in  running  water  for  from 
6  to  24  hours.  From  water  they  are  transferred  to  gradually 
increasing  strengths  of  alcohol.  Particles  of  fat  are  blackened  by 
the  mixture.  Sections  stain  well  with  safranin  or  hematoxylin. 
Read  the  remarks  on  osmic  acid,  21. 
•  13.  Chrom- Ace  tic-Formalin  Mixture. — 

Chromic  acid,  1  per  cent  solution 16  parts 

Glacial  acetic  acid 1  part 

Just  before  using  add  to  two  volumes  of  this  mixture  one  volume 
of  formalin.  This  is  a  good  fixing  fluid  for  general  embryological 
work. 

14.  Corrosive  Sublimate  (mercuric  chloride,  bichloride  of 
mercury). — Corrosive  sublimate  is  ordinarily  used  as  a  saturated 
solution  in  distilled  water  (about  a  7  per  cent  solution)  or  in  normal 
saline.  The  latter  keeps  better  and  contains  a  greater  percentage  of 
the  sublimate.  Corrosive  sublimate  is  an  excellent  and  rapid  fixing 
fluid  for  many  objects  (glands,  epithelia,  etc.).  Objects  should 
remain  in  the  fluid  only  long  enough  to  become  thoroughly  fixed; 
this  has  been  accomplished  when  they  have  become  opaque  through- 
out. Only  a  few  minutes  or  even  seconds  are  required  to  fix  very 
delicate  objects,  but  denser  tissues  may  require  from  4  to  24  hours. 
The  value  of  the  fluid  is  usually  enhanced  by  the  addition  of  5  per 
cent  of  glacial  acetic  acid.  Small  pieces  of  tissue  (not  over  0 . 6  cm. 
in  diameter)  should  be  used  where  practicable.  Washing  may  be 
done  in  running  water  (several  hours)  or  in  50  to  70  per  cent  alcohol. 

CAUTIONS. — (1)  With  corrosive  sublimate  or  mixtures  containing 
it,  the  mercuric  salt  is  often  not  wholly  removed  in  washing.  If  the 
tissues  are  to  remain  several  days  or  weeks  in  alcohol,  the  alcohol  will 
gradually  extract  it.  If  they  are  to  be  used  within  a  few  days, 


Some  Standard  Reagents  and  Their  Uses  213 

however,  it  is  necessary  to  remove  the  excess  of  sublimate  by  adding 
a  few  drops  of  a  10  per  cent  alcoholic  solution  of  iodine  to  the  70  per 
cent  alcohol.  Sufficient  of  the  solution  is  added  to  give  the  alcohol  a 
port-wine  color;  as  often  as  the  color  disappears  the  iodine  must  be 
renewed.  After  from  12  to  48  hours  of  this  treatment,  the  iodine  color 
persists,  and  the  object  should  then  be  transferred  to  fresh  70  or 
80  per  cent  alcohol,  which  must  be  renewed  until  it  no  longer  extracts 
iodine  from  the  specimen.  Some  workers  prefer  not  to  treat  tissues 
fixed  in  a  mercuric  fixer  with  the  iodized  alcohol  until  they  are 
sectioned  and  on  slides.  The  treatment  then  requires  only  about 
30  minutes. 

(2)  In  handling  corrosive  sublimate,  a  glass  or  horn  spoon  should 
be  used  instead  of  a  metal  instrument,  because  it  corrodes  metal. 

(3)  Use  distilled  water,  not  tap  water,  in  making  an  aqueous 
solution. 

15.  Corrosive  Sublimate  and  Acetic  Acid. — 

Corrosive  sublimate,  saturated  aqueous  solu- 
tion            100  parts 

Glacial  acetic  acid 5  to  10  parts 

This  is  an  excellent  reagent  for  embryonic  tissues  and  for  organs 
which  do  not  contain  a  very  great  amount  of  connective  tissue.  See 
remarks  under  14. 

16.  Corrosive  Sublimate,  Nitric- Acid  Mixture  (Gilson's  mercuro- 
nitric  mixture) . — 

Corrosive  sublimate 5  grams 

Nitric  acid  (approximately  80  per  cent) 4  c.c. 

Glacial  acetic  acid 1  c.c. 

Alcohol  (70  per  cent) 25  c.c. 

Distilled  water 220  c.c. 

Filter  after  three  days. 

Gilson's  is  an  excellent  general  reagent  and  gives  a  very  delicate 
fixation.  Objects  should  be  left  in  the  fluid  from  15  to  30  minutes  for 
delicate  ones  to  6  hours  for  those  which  are  larger  or  denser,  although 
many  tissues  may  be  left  for  24  hours  without  injury.  This  is  one 
of  the  most  satisfactory  killing  and  fixing  reagents  that  the  beginner 
can  use. 


214  Animal  Micrology 

Danchakoff's  Mixture.— See  7  B. 
Erlicki's  Fluid.— See  8. 

17.  Ether- Alcohol. — Equal  parts  of  sulphuric  ether  and  absolute 
alcohol. 

Flemming's  Solution. — See  12. 

18.  Formalin. — See  reagent  6,  p.  8,  and  reagent  4,  p.  29.     It 
should  be  borne  in  mind  that  formalin  is  a  reducing  agent  and  will 
rapidly  decompose  such  reagents  as  osmic  acid  or  chromic  acid  if 
mixed  with  them.     It  preserves  fat  and  myelin,  so  that  they  may 
be  stained  by  the  standard  methods,  and  various  substances,  such  as 
amyloid  and  hemosiderin,  to  which  it  may  be  desirable  to  apply 
chemical  tests. 

Commercial  formalin  is  always  slightly  acid.  This  is  not  objec- 
tionable for  ordinary  fixation.  If  neutral  formalin  is  required,  add 
magnesium  carbonate  to  the  commercial  variety,  keeping  a  deposit 
of  the  carbonate  on  the  bottom  of  the  formalin  container. 

19.  Formalin,  Alcohol,  and  Acetic  Acid  (Lavdowsky's  mixture). — • 

Formalin,  commercial 10  parts 

Alcohol,  95  per  cent 50  parts 

Glacial  acetic  acid 2  parts 

Distilled  water 40  parts 

This  mixture  is  recommended  in  some  cases  for  the  treatment  of 
embryos,  especially  when  the  nervous  system  is  to  be  studied.  It 
penetrates  well  and  preserves  faithfully;  the  alcohol  counteracts  the 
swelling  effects  of  the  acetic  acid  and  the  formalin.  Material  may 
remain  in  it  without  injury  for  several  days.  The  fluid  should  sooner 
or  later  be  replaced  by  70  per  cent  alcohol.  No  preliminary  washing 
is  necessary. 

Formalin-Zenker. — See  7. 

20.  Formol  Sublimate  (Worcester's  fluid). — a)  Make  a  saturated 
solution  of  corrosive  sublimate  in  10  per  cent  formalin.     This  reagent 
is  recommended  by  Raymond  Pearl  (Journal  of  Applied  Microscopy, 
VI,  2451)  as  "extremely  satisfactory'7  for  killing  and  fixing  protozoa. 
Washing  may  be  done  in  water  or  4  per  cent  formalin.     The  material 
may  be  preserved  in  4  per  cent  formalin  or  carried  up  the  grades  of 
alcohol  to  70  per  cent  alcohol. 


Some  Standard  Reagents  and  Their  Uses  215 

b)  If  to  9  parts  of  this  formol-sublimate  mixture  1  part  of 
glacial  acetic  acid  is  added,  Worcester's  formol-sublimate-acetic 
mixture  is  obtained.  Pearl  recommends  this  highly  for  teleost  eggs 
and  for  embryological  material  in  general.  It  will  not  produce 
coagulations  and  cloudiness  in  the  gelatinous  envelopes  of  amphibian 
eggs,  if  thoroughly  washed  out  after  fixing.  Preservation  is  the 
same  as  for  (a).  Johnson  (Journal  of  Applied  Microscopy,  VI,  2652) 
also  recommends  this  reagent  very  highly  for  general  work  except 
in  the  case  of  nervous  tissue. 

Personally,  I  have  found  it  advisable  not  to  prepare  either  of  the 
above  mixtures  until  needed  because  the  formalin,  which  is  a  reducing 
agent,  causes  much  of  the  mercuric  salt  to  pass  over  into  the  insoluble 
mercurous  salt. 

Gilson's  Mercuro- Nitric  Mixture. — See  16, 

Kelly's  Fluid.— See  7  A. 

Hermann's  Fluid.— See  27. 

Kleinenberg's  Picro-Sulphuric. — See  26. 

Lavdowsky's  Mixture. — See  19. 

Miiller's  Fluid.— See  9. 

21.  Osmic  Acid  (really  the  tetroxide  of  osmium  Os04). — Osmic 
acid  kills  quickly  and  fixes  well.  It  is  exceedingly  volatile.  The 
chief  objections  to  it,  aside  from  its  extremely  poisonous  nature,  are 
its  poor  powers  of  penetration,  and  the  fact  that  it  becomes  reduced 
in  the  presence  of  the  least  amount  of  dust  containing  organic  par- 
ticles. The  substance  must  be  handled  with  the  greatest  care,  as 
even  the  vapors  are  dangerous.  It  is  usually  put  up  in  small  quanti- 
ties (0.1  to  1  gram)  in  hermetically  sealed  glass  tubes.  In  making 
up  solutions,  the  wrappings  are  removed  from  such  a  tube,  and  the 
tube  is  dropped  into  a  reagent  bottle,  where  it  may  then  be  broken 
by  means  of  a  glass  rod.  Aside  from  its  use  in  mixtures  (see  reagents 
12  and  27),  the  vapor  or  a  0.05  to  a  1  per  cent  aqueous  solution  is 
commonly  used.  A  stock  solution  of  1  per  cent  is  usually  kept  oij 
hand.  It  must  be  kept  free  from  dust.  As  the  most  practical  way 
of  preventing  reduction,  Lee  recommends  that  the  osmic  acid  for 
ordinary  work  be  kept  as  a  solution  in  chromic  acid  (a  2  per  cent 
solution  of  osmic  acid  in  a  1  per  cent  aqueous  solution  of  chromic 


216  ^  nimal  Micrology 

acid).  This  solution  may  be  employed  in  making  up  Flemming's 
solution  or  for  the  purpose  of  fixation  by  means  of  osmium  vapor. 
For  vapor  fixation,  however,  many  workers  prefer  the  vapor  from  the 
solid  crystals. 

To  fix  by  means  of  the  vapor,  the  tissue  is  pinned  to  the  lower 
end  of  a  cork  which  fits  tightly  into  the  bottle  containing  the  osmic 
acid,  or  it  is  suspended  by  a  thread.  Objects  which  will  adhere  to  a 
slide  are  fixed  by  simply  inverting  the  slide  over  the  mouth  of  the 
bottle.  The  time  required  for  such  fixation  varies  from  thirty  seconds 
or  a  few  minutes  for  isolated  cells  to  several  hours  for  thicker  objects, 
such  as  the  retina.  For  fixing  in  the  solution,  24  hours  are  required 
ordinarily.  Objects  are  then  washed  in  running  water  for  the  same 
length  of  time.  Only  small  or  thin  pieces  can  be  fixed  by  means  of 
either  the  solution  or  the  vapor.  The  stains  which  follow  osmic 
acid  best  are  hematoxylin,  methyl  green  (for  study  in  aqueous  media), 
alum-carmine,  picro-carmine,  and  safranin. 

22.  Picric   Acid. — A   cold   saturated   aqueous   solution    (about 
1.2  per  cent)  of  picric  acid  is  commonly  used.     Small  objects  are 
fixed  in  from  a  few  minutes  (infusoria)  to  6  hours;  objects  up  to  1  cm. 
in  size  in  from  24  to  36  hours.     They  may  be  left  a  much  longer  time, 
however,  without  injury.     Large  objects  may  require  weeks  for 
proper  fixation.     After  fixing,  tissues  should  be  washed  in  70  per 
cent  alcohol  until  the  alcohol  is  no  longer  colored  by  the  picric  acid. 
The  tissue  should  not  pass,  during  subsequent  treatment  (with  a 
few  exceptions  in  case  of  staining),  into  an  aqueous  medium  or  into  an 
alcohol  of  less  than  70  per  cent  strength,  because  such  media  seem  to 
undo  the  work  of  fixation. 

23.  Picric  Alcohol. — Gage  recommends  a  0.2  per  cent  solution  of 
picric  acid  in  50  per  cent  alcohol  as  an  excellent  fixer  and  hardener 
for  almost  any  tissue  or  organ.     Time  required,  1  to  3  days.     Entire 
objects  which  have  been  fixed  in  picric  acid  or  in  picric  alcohol  stain 
readily  in  borax-carmine  or  paracarmine. 

24.  Picro-Acetic. — Saturate  a  1  per  cent  aqueous  solution  of  acetic 
acid  with  picric  acid.     This  liquid  is  widely  used  as  a  general  reagent, 
and  is  to  be  preferred  for  most  purposes  to  picric  acid  alone.     For 
washing,  etc.,  see  remarks  under  22. 


Some  Standard  Reagents  and  Their  Uses  217 

Picro- Acetic-Formalin. — See  Bouin's  Picro-Formol,  pp.  9,  29. 

25.  Picro-Sublimate.— 

Rabl's: 

Picric  acid,  saturated  aqueous  solution 1  vol. 

Corrosive  sublimate,  saturated  aqueous  solution  1  vol. 

Distilled  water 2  vols. 

This  mixture  has  been  especially  recommended  for  embryos. 
They  are  left  in  the  fluid  for  12  hours,  then  washed  in  weak  alcohol 
and  transferred  to  gradually  increasing  strengths  of  alcohol. 

0.  vom  Rath's: 

Picric  acid,  cold  saturated  solution 1  vol. 

Corrosive  sublimate,  hot  saturated  solution 1  vol. 

Glacial  acetic  acid 0.5  to  1  vol. 

After  fixing  for  several  hours,  transfer  the  material  directly  into 
alcohol. 

26.  Picro-Sulphuric  (Kleinenberg's). — 

Picric  acid,  saturated  aqueous  solution 98  vols. 

Sulphuric  acid 2  vols. 

Water 200  vols. 

This  is  an  excellent  reagent  for  embryos,  either  for  entire  mounts 
or  for  sectioning.  Chick  embryos  of  24  to  48  hours  should  remain 
in  the  liquid  for  from  2  to  4  hours;  older  embryos  for  from  3  to  6 
hours.  For  washing,  70  per  cent  alcohol  is  used.  It  should  be 
changed  (frequently  at  first)  until  the  color  ceases  to  come  out  of 
the  embryos.  Preserve  in  about  80  per  cent  alcohol. 

Lillie  recommends  the  addition  of  glacial  acetic  acid  sufficient 
to  make  a  5  per  cent  solution  of  acetic  acid. 

27.  Platino-Aceto-Osmic  Mixture  (Hermann's  fluid). — 

Platinum  chloride,  1  per  cent  aqueous  solution. .  .    60  c.c. 

Osmic  acid,  2  per  cent  aqueous  solution 8  c.c. 

Glacial  acetic  acid 4  c.c. 

Hermann's  fluid  is  one  of  the  most  valuable  cytological  reagents. 
Only  small  pieces  of  tissue  should  be  used.  The  washing  and  sub- 
sequent treatment  are  the  same  as  for  Flemming's  solution  (12). 


218  Animal  Micrology 

For  subsequent  treatment  with  pyrogallol,   see  70.     Read,   also, 
remarks  on  osmic  acid  (21). 

RabPs  Picro-Sublimate.-^-See  25. 

Rath's  (O.  vom)  Picro-Sublimate.— See  25. 

Ripart  and  Petit,  Liquid  of. — See  10. 

Tellyesnicky's  Fluid.— See  5. 

Van  Gehuchten's  Fluid. — Same  as  2,  6). 

Worcester's  Fluid.— See  20. 

Zenker's  Fluid.— See  6. 

II.    STAINS 

Read  the  general  statement  about  stains  in  chap.  ii. 

28.  Alum-Cochineal. — For  formula  see  p.  9.     Alum-cochineal  is 
one  of  the  best  stains  for  entire  objects.     It  is  easy  to  work  with, 
and  does  not  over  stain.     The  time  required  for  staining  is  from  24  to 
36  hours  ordinarily.     After  staining,  the  object  should  be  washed  in 
water  for  15  or  20  minutes  to  extract  the  alum,  which  would  otherwise 
crystallize  when  the  preparation  is  placed  in  alcohol.     Too  long  an 
immersion  in  water  may  extract  the  stain  to  too  great  an  extent. 
From  water  the  object  should  be  passed  upward  through  the  grades 
of  alcohol,  remaining  about  an  hour  in  each.     The  writer  has  found 
alum-cochineal  especially  valuable  for  flatworms  (tapeworms,  flukes, 
etc.)  and  enfbryos.     If  it  is  desired  to  use  a  counterstain  with  it, 
Lyon's  blue,  picric  acid,  orange  G,  or  light  green  will  answer. 

29.  Alum-Carmine. — 

Powdered  carmine 1  gram 

Ammonia  alum  (2 . 5  per  cent  aqueous  solution)   100  c.c. 

Boil  f  oft|  minutes,  and  filter  when  cool.  The  uses  and  manipula- 
tion are  the  same  as  for  reagent  28.  These  stains  affect  calcareous 
structures  injuriously. 

30.  Anilin  Stains. — -Read  the  general  remarks  about  anilin  stains 
in  chap,  ii  (p.  20).     The  formulae  for  some  of  the  most  important  are 
given  separately  in  this  list  in  their  proper  alphabetical  position. 

The  dyes  are  dissolved  in  water,  in  alcohol  of  any  desired  strength, 
or  in  anilin  water,  according  as  they  are  soluble  in  these  media,  or  as 
they  meet  the  needs  of  the  operator.  Some  workers  even  use  some 


Some  Standard  Reagents  and  Their  Uses  219 

of  them  as  counterstains  dissolved  in  the  clearing  fluid.  For  the 
study  of  nuclei,  after  Hermann's  or  Flemming's  fluid  has  been  used 
for  fixing,  the  writer  has  found  a  weakly  alcoholic  anilin-water  solu- 
tion to  be  the  most  satisfactory.  As  cytoplasmic  contrast  stains 
alcoholic  solutions  (in  70  to  95  per  cent  alcohol)  have  given  the  best 
results.  Anilin  water  is  made  by  shaking  up  4  c.c.  of  anilin  oil  in 
90  c.c.  of  distilled  water  and  filtering  the  mixture  through  a  wet  filter. 
Enough  alcohol  may  be  added  to  make  it  a  20  per  cent  alcohol,  if  a 
weakly  alcoholic  solution  is  desired. 

The  length  of  time  which  sections  should  be  immersed  in  the 
stain  varies  from  a  few  seconds  or  minutes  for  some  of  the  dyes 
(especially  when  .used  for  cytoplasm)  to  24  to  36  hours  for  others 
(especially  nuclear).  Sections  usually  overstain,  in  which  case 
they  are  differentiated  by  means  of  alcohol,  either  pure  or  slightly 
acidulated  with  hydrochloric  acid.  The  color  is  thus  extracted 
rapidly;  decolorization  should  be  stopped  immediately  after  the 
color  ceases  to  come  from  the  tissue  in  clouds  (20  seconds  to  3  min- 
utes) .  If  acidulated  alcohol  is  employed,  it  must  be  in  much  weaker 
solution  than  that  used  for  extracting  carmines  or  heft&atoxylins. 
One  part  of  hydrochloric  acid  to  1,000  of  water  or  alcohol  is  about  the 
correct  proportion.  When  one  desires  to  study  the  karyokinetic 
figures  of  nuclei,  the  acid-alcohol  differentiation  should  be  employed, 
but  if  resting  nuclei  are  to  be  studied,  only  neutral  alcohol  should 
be  used. 

31.  Anilin  Blue,  Orange  G,  and  Acid  Fuchsin  (Mallory's  triple 
connective-tissue  stain) . — • 

Solution  I: 

Acid  f uchsin 0.5 

Distilled  water 100     c.c. 

Solution  II: 

Anilin  blue  (Griibler's  water  soluble) 0.5  gram 

Orange  G  (Griibler) 2.0  grams 

Phosphomolybdic  acid,  1  per  cent  aqueous 

solution 100      c.c. 

The  tissue  should  have  been  fixed  in  Zenker's  fluid.  Stain 
either  paraffin  or  celloidin  sections  in  the  acid-fuchsin  solution  for 


220  Animal  Micrology 

5  minutes  or  longer,  depending  upon  the  freshness  of  the  tissue. 
Transfer  directly  to  solution  II  and  stain  for  from  10  to  20  minutes 
or  longer.  Wash  and  dehydrate  in  several  changes  of  95  per  cent 
alcohol.  Pass  paraffin  sections  through  absolute  alcohol,  clear  in 
xylol,  and  mount  in  balsam.  Clear  celloidin  sections  from  95  per 
cent  alcohol  in  creosote  or  other  celloidin  clearer,  or  blot  and  clear 
in  xylol,  finally  mounting  in  balsam. 

Connective-tissue  reticulum,  collagen  fibrils,  mucus,  amyloid, 
and  various  other  hyaline  substances  stain  in  different  shades  of 
blue;  nuclei,  cytoplasm,  axis-cylinders,  neuroglia  fibers,  fibroglia  fib- 
rils, and  fibrin  stain  red ;  elastic  fibers,  pale  pink  or  yellow ;  and  red 
blood  corpuscles  and  myelin  sheaths,  yellow.  If  the  acid  fuchsin 
is  omitted,  nuclei  and  protoplasm  stain  yellow  and  the  connective- 
tissue  fibrillae  and  reticulum  stand  out  sharply  in  deep  blue.  This 
is  an  excellent  stain  for  developing  bone,  inasmuch  as  cartilage 
stains  light  blue  and  bone  dark  blue. 

Bensley's  Acid  Fuchsin-Methyl  Green  Method. — See  p.  144. 

Bensley's  Copper  Chrome  Hematoxylin  Method. — See  p.  145. 

32.  Bismarck  Brown.— Boil  1  gram  of  the  stain  in  100  c.c.  of 
water,  filter,  and  add  30  c.c.  of  strong  alcohol.     Bismarck  brown  is 
a  nuclear  stain  which  does  not  overstain,  although  it  acts  rapidly. 
After  staining,  wash  in  95  per  cent  or  absolute  alcohol.     This  stain 
is  also  used  in  aqueous  solution  for  intra-vitam  staining;  the  nucleus 
of  the  living  cell  may  thus  be  colored.     It  has  been  used  as  an  intra- 
vitam  stain  mostly  in  the  study  of  infusoria.     The  stain  may  be 
fixed  by  means  of  a  0.2  per  cent  chromic-acid  solution,  but  this,  of 
course,  destroys  the  life  of  the  cells. 

33.  Borax-Carmine  (Grenadier's). — 

Borax  (4  per  cent  aqueous  solution) 100  c.c. 

Carmine 1  gram 

Boil  until  the  carmine  dissolves,  then  add  100  c.c.  of  70  per  cent 
alcohol.  Filter  after  24  hours. 

This  is  a  stain  much  used  in  the  past  for  staining  in  bulk.  Objects 
must  be  left  in  it  for  from  24  hours  to  several  days.  They  are  then 
transferred,  without  washing,  to  acid  alcohol  and  left  until  the  color 
no  longer  comes  away  in  clouds.  Objects  should  become  bright 


Some  Standard  Reagents  and  Their  Uses  221 

scarlet  in  color.     Finally  they  should  be  washed  and  hardened  in 
neutral  alcohol. 

34.  Bordeaux  Red.— 

Bordeaux  red 1  gram 

Distilled  water : 100  c.c. 

This  is  a  good  plasma  stain.  Recommended  by  Heidenhain 
as  a  contrast  stain  for  iron-hematoxylin  in  the  demonstration  of 
centrosomes  (p.  147).  Stain  for  12  to  24  hours. 

35.  Carmalum  (Mayer's). — 

Carminic  acid 1  gram 

Alum 10  grams 

Distilled  water , 200  c.c. 

Dissolve  with  heat  and  filter  the  solution  when  cold.  Add  a 
few  crystals  of  thymol  or  a  little  salicylic  acid  to  prevent  the  forma- 
tion of  mold.  Carmalum  is  one  of  the  best  stains  for  staining  objects 
in  bulk  and  will  follow  almost  any  fixing  reagent,  even  osmic  acid. 
If  the  object  has  an  alkaline  reaction  it  does  not  stain  so  well.  Wash- 
ing is  done  in  water. 

36.  Carmine  (Scale's).— 

Powdered  carmine .  .  .- 1  gram 

Ammonia 3  C.C. 

Pure  glycerin 96  c.c. 

Distilled  water 96  c.c. 

Alcohol,  95  per  cent 24  c.c. 

The  ammonia  and  part  of  the  water  are  first  mixed  and  the 
carmine  dissolved  in  the  mixture.  The  remaining  water  is  added 
and  the  solution  is  left  in  an  open  dish  until  the  ammonia  has  almost 
evaporated.  The  alcohol  and  glycerin  are  then  added.  For  staining, 
equal  parts  of  the  stain  and  glycerin  are  used.  The  staining  is  carried 
on  for  24  hours  under  a  bell-jar  in  an  uncovered  dish.  A  second  open 
dish  containing  acetic  acid  is  placed  under  the  bell-jar.  After 
staining,  the  sections  are  washed  in  water,  then  in  weak  hydro- 
chloric acid  (1  to  500  of  water),  and  again  in  water.  Minot  recom- 
mends this  stain  and  method  of  treatment  especially  for  the  placenta 
and  for  the  central  nervous  system  of  embryos. 


222  Animal  Micrology 

37.  Carmine,  Picric  Acid,  and  Indigo  Carmine  (Calleja's  staining 
fluid).— 

Solution  I: 

Carmine 2        grams 

Lithium  carbonate,  saturated  aqueous  solu- 
tion      100        c.c. 

Solution  II: 

Indigo-carmine 0 . 25  gram 

Picric  acid,  saturated  aqueous  solution.  ...    100        c.c. 

Place  sections  in  solution  I  for  from  5  to  10  minutes,  then  into 
acid  alcohol  until  they  become  pale  red  (20  to  30  seconds) ;  wash  well 
in  water.  Next  place  the  sections  in  solution  II  for  5  to  10  min- 
utes, then  into  acetic  acid  (0.2  to  0.5  per  cent)  for  a  few  seconds, 
and  wash  well  in  water.  Dehydrate  rapidly  and  clear  in  xylol. 
The  method  is  useful  for  epithelial  cells  and  connective  tissue. 

38.  Carmine,  Acid  (Schneider's). — Add  carmine  to  boiling  acetic 
acid  of  45  per  cent  strength  until  no  more  will  dissolve.     Filter  the 
solution  when  cool.     This  is  a  valuable  reagent  for  the  study  of  the 
nuclei  of  fresh  cells.     It  is  very  penetrating  and  gives  a  brilliant 
stain.     The  strong  acetic  acid  ultimately  destroys  the  cell. 

39.  Congo  Red. — For  formula,  see  p.  10.     This  is  a  good  counter- 
stain  for  hematoxylin,  especially  when  applied  to  fetal  and  young 
tissues.     Its  solutions  become  blue  in  the  presence  of  free  acid,  hence 
it  is  useful  in  determining  the  existence  of  free  acid  in  tissues. 

40.  Cyanin  (Chinolin  Blue;   Quinoline  Blue). — Dissolve  1  gram 
of  cyanin  (prepared  by  H.  A.  Metz  &  Co.,  of  New  York)  in  100  c.c.  of 
95  per  cent  alcohol,  and  add  100  c.c.  of  distilled  water.     This  is  a  good 
cytological  stain.     Sections  are  stained  for  5  to  10  minutes.     Chromo- 
somes stain  a  deep  blue.     I  have  found  cyanin  followed  by  erythrosin 
(0 . 5  per  cent  alcoholic  solution)  especially  valuable  for  spermatozoa. 

41.  Ehrlich-Biondi  Triple  Stain  (Heidenhain) .— The  ingredients 
should  be  obtained  from  Griibler  and  Hollborn,  Baiersche  Strasse  63, 
Leipzig,  or  from  their  agents. 

Acid  f uchsin,  saturated  aqueous  solution 4  parts 

Orange  G  "  "       7  parts 

Methyl  green  (Methylgrun  00)  saturated  aqueous 

solution 8  parts 


Some  Standard  Reagents  and  Their  Uses  223 

The  solution  of  orange  should  be  prepared  first,  and  the  solutions 
of  fuchsin  and  methyl  green  added  to  it  with  continual  stirring. 
Each  solution  must  be  thoroughly  saturated;  it  takes  several  days  for 
this  to  occur.  The  above-mentioned  mixture  constitutes  a  stock  solu- 
tion which  should  be  diluted  with  about  50  or  100  times  its  volume  of 
water  before  using.  According  to  Lee  (Microtomisf  s  Vade-Mecum), 
"if  a  drop  be  placed -on  blotting  paper  it  should  form  a  spot  bluish 
green  in  the  center,  orange  at  the  periphery.  If  the  orange  zone  is 
surrounded  by  a  broader  red  zone,  the  mixture  contains  too  much 
fuchsin."  For  use  with  this  method,  tissues  should  be  fixed  in  pure 
corrosive-sublimate  solution.  Sections  should  be  thin  (3  to  5  microns) 
and  must  remain  in  the  stain  from  12  to  24  hours.  They  should  then 
be  rapidly  washed  in  95  per  cent  alcohol,  placed  for  a  short  time  in 
absolute  alcohol,  and  cleared  in  xylol.  If  the  sections  remain  in 
the  alcohols  any  considerable  length  of  time,  the  methyl  green  will 
be  extracted.  The  stain  is  very  uncertain  in  its  action,  but  when  it  is 
successfully  applied  the  results  are  excellent.  It  is  used  chiefly  in 
cytological  studies,  especially  in  connection  with  gland  cells.  Griibler 
prepares  a  dry  powder  for  this  three-color  mixture,  but  the  results  are 
usually  not  as  satisfactory  as  when  the  mixture  is  properly  made 
fresh.  To  prepare  the  stain  from  the  powder,  a  0 . 4  per  cent  solution 
of  the  latter  in  distilled  water  is  made,  and  to  100  c.c.  of  this  solution 
7  c.c.  of  a  0. 5  per  cent  aqueous  solution  of  acid  fuchsin  is  added. 

42.  Ehrlich's  Triple  Stain.— For  blood  films  Ehrlich's  so-called 
triacid  mixture  is  a  serviceable  stain  which  is  widely  used. 

Orange  G,  saturated  aqueous  solution 14  c.c. 

Acid  fuchsin,  saturated  aqueous  solution 7  c.c. 

Distilled  water 15  c.c. 

Absolute  alcohol 25  c.c. 

Methyl  green,  saturated  aqueous  solution 12  c.c. 

Glycerin 10  c.c. 

Each  solution  must  be  thoroughly  saturated  (several  days) .  Add 
the  ingredients  in  the  order  named,  shaking  the  mixture  well  before 
each  addition.  It  is  best  for  the  stain  to  stand  several  weeks  before 
it  is  used.  Neutrophil  granules  stain  violet,  oxyphil  granules  a 
brownish  red.  The  mixture  stains  in  from  5  to  15  minutes. 


224  Animal  Micrology 

43.  Eosin. — See  reagent  16,  p.  10.     This  anilin  dye  is  often  used 
after  hematoxylin  as  a  contrast  stain.     It  is  specific  for  certain 
granules  of  leucocytes  and  for  red  blood  corpuscles,  giving  to  the 
latter  a  very  characteristic  coppery-red  tinge.     Some  workers  prefer 
to  dissolve  it  in  water  or  in  some  cases  in  the  clearer. 

44.  Erythrosin. — An  eosin;   properties  and  manipulation  much 
the  same  as  ordinary  eosin  (see  reagent  43). 

45.  Fuchsin,  Acid  (Rubin  S,  Acid  Magenta,  Magenta  S). — 

Acid  f  uchsin 0.5  gram 

Distilled  water 100      c.c. 

/^This  is  an  excellent  anilin  stain  for  cytoplasmic  structures.  It 
is  also  used  in  some  instances  as  a  specific  stain  for  nerve  tissue. 
Acid  fuchsin  should  not  be  confounded  with  basic  f  uchsin,  which  is  a 
nuclear  stain.  It  too  is  used  in  aqueous  solution.  When  fuchsin 
alone  is  mentioned  by  writers,  without  specifying  whether  it  is  acid 
or  basic,  the  basic  fuchsin  is  ordinarily  meant. 

46.  Fuchsin-Methyl-Green  Stain  (Auerbach's).— Keep  in  sepa- 
rate bottles  0 . 1  per  cent  aqueous  solutions  of  acid  fuchsin  and  methyl 
green  respectively.    When  ready  to  use,  mix  2  parts  of  the  acid- 
fuchsin  solutfon  with  3  or  4  parts  of  the  methyl  green,  after  acidulating 
every  50  c.c.  of  the  former  with  1  drop  of  a  10  per  cent  solution  of 
acetic  acid. 

This  stain  works  best  after  a  sublimate  fixer.  Chromosomes 
stain  green,  linin  and  plasmosomes  red.  Sections  should  not  be  over 
3  or  4  microns  thick.  Stain  for  15  minutes  and  transfer  directly 
to  95  per  cent  alcohol.  As  soon  as  the  green  stain  ceases  to  leave 
the  sections  in  clouds,  pass  the  slides  rapidly  through  absolute 
alcohol  and  xylol  and  mount  in  balsam. 

47.  Fuchsin  (Acid)  and  Picric  Acid  (Van  Giesen's  stain). — 

Acid  fuchsin,  1  per  cent  aqueous  solution 10  c.c. 

Picric  acid,  saturated  aqueous  solution 90  c.c. 

This  stain  is  frequently  used  in  conjunction  with  hematoxylin 
in  the  study  of  fibrous  or  of  nerve  tissue.  Small  bits  of  tissue  should 
be  fixed  in  corrosive  sublimate  or  its  mixtures.  Sections  are  slightly 
overstained  with  hematoxylin,  rinsed  in  water,  and  then  stained 


Some  Standard  Reagents  and  Their  Uses  225 

5  minutes  in  the  picro-fuchsin  mixture.  To  avoid  extracting  toe 
much  of  the  yellow  color  in  dehydrating  and  clearing,  the  alcohols  and 
clearer  should  each  have  a  few  crystals  of  picric  acid  added  to  them. 
The  result  should  be:  nuclei  and  epithelia  brown;  white  fibrous 
connective  tissue  red;  elastic  tissue  and  muscle  yellow. 

48.  Gentian  Violet. — This  is  one  of  the  best  of  the  nuclear  anilin 
stains.     It  is  best  made  up  in  anilin  water  and  weak  alcohol  (see 
reagent  30). 

Gentian  violet 1  gram 

Anilin  water 80  c.c. 

Alcohol,  95  per  cent 20  c.c. 

The  stain  works  well  with  thin  sections.  It  is  also  widely  used 
in  the  study  of  bacteria.  For  differentiation,  Gram's  method  is  used. 

Gram's  solution: 

Iodine 1  gram 

Iodide  of  potassium 2  grams 

Water 300  c.c. 

After  staining,  the  sections  are  placed  in  this  solution  until  they 
are  black  (2  to  3  minutes)  and  are  then  decolorized  in  absolute  alcohol 
until  they  appear  gray.  See  also  reagent  73. 

49.  Gold  Chloride. — The  gold-chloride  method  is  used  chiefly 
in  the  study  of  nerve-fiber  terminations,  both  motor  and  sensory, 
although  it  is  sometimes  used  for  the  coloration  of  other  tissue  ele- 
ments (capsules  of  cartilage,  etc.) .    The  process  is  really  an  impregna- 
tion; through  the  agency  of  sunlight  and  of  certain  reagents  (acetic, 
citric,  formic,  or  oxalic  acid)  the  gold  is  deposited  in  the  tissues  in  the 
form  of  very  fine  particles.     There  are  numerous  modifications  of  the 
method,  one  of  which  is  given  in  chap.  ix. 

50.  Golgi's  Chrome-Silver  Method. — See  chap.  ix.  p.  71. 
Hematoxylin. — For  general  statement  see  chap,  ii,  p.  20,  and  the 

remarks  under  12,  p.  9. 

51.  Hematoxylin,  Conklin's  Picro. — • 

Delafield's  hematoxylin 1  part 

Water 4  parts 


226  Animal  Micrology 

Add  one  drop  of  Kleinenberg's  picro-sulphuric  (26)  to  each 
cubic  centimeter  of  the  solution.  This  is  a  good  stain  (1  to  3  hours) 
for  embryos  which  are  to  be  mounted  entire.  If  the  embryos  are  to 
be  sectioned  they  should  be  stained  for  12  hours.  ' 

52.  Hematoxylin,  Delafield's. — See  reagent  12,  p.  9. 

53.  Hematoxylin,  Ehrlich's  Acid. — • 

Hematoxylin 2  grams 

Absolute  alcohol 100  c.c. 

Glacial  acetic  acid 10  c.c. 

Glycerin 100  c.c. 

Distilled  water 100  c.c. 

Potassium  alum 10  grams 

Dissolve  the  hematoxylin  in  the  acetic  acid  with  25  c.c.  of  the 
alcohol;  then  add  the  glycerin  and  the  remaining  alcohol.  Dissolve 
the  alum  in  the  water  by  the  aid  of  heat  and  slowly  pour  the  warm 
solution  while  stirring  into  the  solution  of  hematoxylin.  The 
solution  must  be  exposed  to  light  and  air  at  least  3  weeks  to  ripen. 
It  is  not  ready  for  use  until  it  acquires  a  deep  red  color.  This  solu- 
tion is  an  excellent  nuclear  stain  and  will  keep  for  years. 

Mann's  acid  hematein  is  the  same  as  this,  except  that  hematein 
(Griibler's)  is  substituted  for  the  hematoxylin.  This  solution  should 
stain  without  having  to  ripen. 

54.  Hematoxylin,  Heidenhain's  Iron-. — See  reagent  17,  p.  11, 
for  formula;    p.  51  for  method;    chap,  xvii  for  cytological  uses. 
This  stain  is  much  used  in  the  study  of  cell  structures  such  as  centro- 
somes,  chromosomes,  etc.     Tissues  are  best  fixed  in  Bouin's,  in  some 
of  the  sublimate  solutions,  or  in  acetic  alcohol,  although  it  will  follow 
liquid  of  Flemming  or  Hermann.     Sections  should  not  be  over  6 
microns  thick.     The  ferric  solution  must  be  renewed  occasionally 
as  it  soon  spoils. 

55.  Hematoxylin,  Mallory's  Phosphotungstic  Acid. — 

Hematein  ammonium 0.1  gram 

Water 100      c.c. 

Phosphotungstic  acid  crystals  (Merck) 2      grams 

Dissolve  the  hematein  by  heating  it  in  a  little  water.  When 
cool  add  it  to  the  rest  of  the  mixture.  If  the  stain  is  too  weak  at 


Some  Standard  Reagents  and  Their  Uses  227 

first,  it  may  be  ripened  by  adding  5  c.c.  of  a  0.25  per  cent  aqueous 
solution  of  permanganate  of  potassium  or  by  allowing  it  to  stand 
two  or  three  weeks.  Hematoxylin  may  be  used  instead  of  hematein 
ammonium  if  10  c.c.  of  the  potassium-permanganate  solution  is 
added  to  ripen  it. 

The  stain  is  recommended  for  the  demonstration  of  neuroglia, 
myoglia,  and  fibroglia  fibrils,  for  fibrin,  and  for  centrosomes  and 
spindles  of  mitotic  figures. 

Janus  Green. — See  pp.  145,  146. 

56.  Light  Green  (Lichtgriin   S.F.).— This  is  a  beautiful  cyto- 
plasmic  anilin  stain  which  is  frequently  used  after  safranin  as  a 
counterstain.     Not  more  than  0 . 5  per  cent  solution  should  be  used 
as  it  stains  very  rapidly  and  very  deeply.     It  may  be  used  either  as  an 
aqueous  or  as  an  alcoholic  solution.    The  writer  has  found  a  0 . 5  per 
cent  solution  in  95  per  cent  alcohol  very  satisfactory.     Sections 
should  remain  in  it  only  a  few  seconds.     Do  not  confuse  it  with 
methyl  green,  which  is  sometimes  called  light  green  by  dealers. 

57.  Lugol's  Solution. — A  solution  of  iodine  in  water  containing 
iodide  of  potassium.     It  is  used  in  various  strengths.    One  of  the 
commonest  formulae  is  that  of  Gram  (p.  225),  although  some  prefer 
a  solution  with  only  one-third  the  amount  of  water  used  by  him. 

58.  Lyons  Blue  (Bleu  de  Lyon). — This  is  one  of  the  best  of  the 
numerous  anilin  blues.     It  is  a  good  contrast  stain  when  used  after 
such  nuclear  stains  as  safranin  and  carmine.    See  reagent  15,  p.  10; 
also  p.  50. 

Magenta,  Acid. — See  45. 

Mallory's  Triple  Connective-Tissue  Stain.— See  31. 

59.  Methylen  Blue. — This  reagent  is  an  extremely  useful  one; 
it  is  of  great  value  in  the  study  of  the  nervous  system,  and  it  can  be 
made  to  give  results  with  intercellular  cement  substance,  lymph 
spaces,  etc.,  as  satisfactory  and  with  greater  certainty  than  impreg- 
nations obtained  with  gold  chloride  or  silver  nitrate.     It  is  also 
serviceable  as  an  intra-vitam  stain.     Furthermore,  methylen  blue 
(saturated  solution  in  70  per  cent  alcohol)  followed  by  eosin  is  some- 
times used  for  the  double  staining  of  blood  corpuscles.     Methylen 
blue  should  not  be  confounded  with  methyl  blue. 


228  Animal  Micrology 

Ordinary  commercial  methylen  blue  usually  contains,  in  addi- 
tion to  the  blue  dye,  a  small  quantity  of  a  reddish-violet  dye.  Such 
methylen  blue  is  termed  polychromatic  and  is  especially  serviceable 
in  staining  certain  cell  granules.  Only  the  pure  methylen  blue, 
however,  should  be  used  for  nerve  staining  and  other  intra-vitam 
work. 

a)  "Intra-Vitam"  Stain  for  Small,  Comparatively  Transparent 
Aquatic  Organisms.— Add  sufficient  methylen  blue  to  the  water 
containing  the  organisms  to  tinge  it  a  light  blue.  Different  tissues 
will  take  up  the  color  after  different  intervals  of  time.  A  given 
tissue  after  having  attained  a  maximum  degree  of  coloration  will 
rapidly  lose  its  color  again.  It  is  necessary,  therefore,  to  watch 
the  organisms  closely  for  the  maximum  of  color  in  the  tissue  desired. 
If  the  observer  wishes,  the  stain  may  be  fixed  for  more  prolonged 
study  by  following  the  processes  indicated  under  (6) .  The  order  in 
which  various  tissues  take  the  stain  seems  to  vary  in  different  organ- 
isms. Usually  gland  cells  stain  first,  then  with  more  or  less  deviation, 
other  epithelial  cells,  fat  cells,  blood  and  lymph  cells,  elastic  fibers, 
smooth  muscle,  and  striated  muscle.  Nerve  cells  and  nerve  fibers 
do  not  ordinarily  take  the  stain  when  the  entire  animal  is  immersed. 

6)  Ehrlich's  Method  for  Nerve-Terminations  and  the  Relations 
of  Nerve  Cells  and  Fibers  to  the  Central  Nervous  System. — The 
stain  should  be  Griibler's  methylen  blue  (rectificiert  nach  Ehrlich). 
A  1  per  cent  solution  in  normal  saline  is  used.  Warm  the  solution 
till  it  steams,  stir  it  thoroughly,  and,  when  cool,  filter.  The  tissue 
must  be  perfectly  fresh.  Chloroform  the  animal  and  immediately 
inject  the  stain  into  the  main  artery  of  the  part  to  be  investigated.  If 
the  animal  is  small,  the  entire  body  may  be  injected.  The  vessels 
should  be  filled  full,  but  care  must  be  taken  not  to  rupture  them.  The 
part  should  become  decidedly  blue  in  color.  It  is  well  after  10  or 
15  minutes  to  inject  more  stain.  At  the  expiration  of  half  an  hour 
after  the  second  injection  remove  small  pieces  of  tissue  containing 
the  nerve  elements  desired,  and  expose  them  freely  to  the  air  on  a  slide 
wet  with  normal  saline.  Examine  every  two  minutes  under  the 
microscope  (without  cover-glass)  until  the  particular  element  to  be 
investigated  (cell,  axone,  termination)  has  developed  a  well-marked 


Some  Standard  Reagents  and  Their  Uses  229^ 

blue  color.  It  is  important  to  catch  the  color  at  the  proper  stage 
and  fix  it  because  it  soon  begins  to  fade. 

Filing  the  Stain. — When  the  desired  element  has  developed  a 
satisfactory  blue  color,  the  tissue  is  transferred  immediately  to  a 
saturated  aqueous  solution  of  Ammonium  pier  ate  (Dogiel's  method) 
and  left  for  from  6  to  24  hours.  For  final  mounting  the  tissue  should 
be  teased  out  sufficiently  to  show  the  proper  elements  and  then 
mounted  in  a  few  drops  6f  a  mixture  of  pure  glycerin  (free  from  acid) 
and  ammonium  picrate  (saturated  aqueous  solution),  equal  parts.  It 
is  well  to  let  the  tissue  stand  in  20  to  30  volumes  of  this  glycerin- 
picrate  mixture  for  a  day  or  two  before  mounting  it.  If  the  prepara- 
tion is  to  be  kept  the  cover-glass  should  be  sealed  (p.  95). 

Sections. — If  it  is  desired  to  make  paraffin  sections  and  mount 
them  in  balsam,  after  treatment  with  the  ammonium  picrate  (10 
to  15  minutes),  the  tissue  must  be  placed  into  20  or  30  volumes  of 
Bethe's  fluid,  which  renders  the  color  insoluble  in  alcohol. 

Bathe's  fluid: 

Molybdate  of  ammonia 1  gram 

Chromic  acid,  2  per  cent  aqueous  solution 10  c.c. 

Hydrochloric  acid,  concentrated  C.P 1  drop 

Distilled  water 10  c.c. 

The  tissue  is  left  in  this  mixture  for  from  45  to  60  minutes  (for 
small  objects)  and  then  washed  1  to  2  hours  in  distilled  water. 
Dehydrate  directly  in  absolute  alcohol;  follow  this  with  xylol, 
imbed  in  paraffin,  and  section  in  the  ordinary  manner.  Sections 
may  be  counterstained  in  alum-carmine  or  alum-cochineal. 

c)  Immersion  Method. — Material  which  cannot  be  readily 
injected  or  which  has  failed  to  stain  may  'be  stained  by  immersion. 
A  0. 1  per  cent  solution  of  the  stain  is  used  (dilute  1  volume  of  the 
solution  used  for  injection  with  9  volumes  of  normal  saline).  To 
small  pieces  (2  to  3  mm.  thick)  of  the  tissue  add  a  few  drops  of  the 
stain  at  intervals  of  about  three  minutes.  The  tissue  should  alwaj^s 
be  moist,  but  never  covered  sufficiently  by  the  solution  to  exclude  air. 
Examine  the  preparation  from  time  to  time  under  the  microscope  and 
when  the  nerve  elements  are  well  stained,  fix  in  ammonium  picrate 
and  proceed  as  in  (6).  In  case  of  the  central  nervous  system,  fairly 


230  Animal  Micrology 

good  results  may  sometimes  be  obtained  by  dusting  the  methylen-blue 
powder  over  the  freshly  cut  surface  of  the  part  to  be  studied.  The 
development  and  fixing  of  the  color  is  the  same  as  in  (6). 

d)  NissPs  Method  of  Staining  Basophil  (Tigroid)  Substance  in 
Nerve  Cells.— 

Methylen  blue 3 . 75  grams 

Venetian  soap  (white  castile  soap) 1 . 75  grams 

Water .....:.   1,000        c.c. 

It  is  best  to  keep  the  stain  for  some  months  before  using. 

Ganglia  should  be  fixed  in  alcohol,  formalin,  or  corrosive  sublimate 
and  sectioned  in  paraffin.  Fix  the  sections  to  the  slide,  dissolve  out 
the  paraffin  with  xylol,  and  run  the  preparation  down  to  the  aqueous 
stain  in  the  ordinary  way.  In  a  test-tube  heat  a  few  cubic  centi- 
meters of  the  stain  until  it  steams,  then  apply  it  while  still  warm  to  the 
sections  on  the  slide,  which  has  been  placed  flat  on  the  desk.  It  takes 
about  6  minutes  for  the  stain  to  act.  Pour  off  the  surplus  stain  and 
rinse  the  slide  in  distilled  water.  Lay  it  flat  on  the  desk  again  and 
flood  the  sections  with  anilin-alcohol  (95  per  cent  alcohol,  9  parts; 
anilin  oil,  1  part).  Let  the  sections  decolorize  (20  to  30  seconds) 
until  they  are  a  pale  blue;  then  drain  off  the  anilin-alcohol  and  trans- 
fer the  preparation  to  absolute  alcohol.  Clear  in  xylol  and  mount 
in  balsam.  The  basophil  granules  should  appear  deep  blue  in  color. 
They  are  arranged  for  the  most  part  concentrically  around  the 
nucleus. 

e)  Unna's  Method  of  Staining  Unstriated  Muscle  in  Sections. — 
Stain  in  a  1  per  cent  aqueous  solution  of  polychromatic  methylen 
blue,  rinse  in  water,  and  then  leave  for  10  minutes  in  a  1  per  cent 
aqueous  solution  of  potassium  ferricyanide.     Transfer  to  acid  alcohol 
until  sufficiently  decolorized,  then  complete  the  dehydration,  and 
mount  in  the  usual  way.     . 

/)  For  Ordinary  Section  Staining  where  a  nuclear  stain  is  desired 
methylen  blue  answers  very  well.  It  is  usually  used  (2  to  24  hours) 
in  aqueous  solution.  The  treatment  is  the  same  as  for  safranin. 

gf)  Impregnation  of  Epithelia,  etc. — Place  the  fresh  tissue,  pref- 
erably a  thin  membrane,  into  a  4  per  cent  solution  of  methylen 
blue  in  normal  saline.  To  demonstrate  the  outline  of  cells,  leave  the 


Some  Standard  Reagents  and  Their  Uses  231 

tissue  in  the  stain  not  longer  than  10  minutes.  To  get  a  negative 
image  of  lymph  spaces,  canals,  etc.,  in  contrast  to  the  ground  sub- 
stance which  becomes  deeply  impregnated,  leave  the  tissue  in  the 
stain  20  to  30  minutes.  For  this  purpose  it  is  advisable  to  remove  any 
membranous  covering  which  invests  the  organ.  In  either  case,  after 
staining,  fix  the  tissue  for  30  to  40  minutes  in  a  saturated  aqueous 
solution  of  ammonium  picrate,  changing  it  once  or  twice,  and  exam- 
ine in  dilute  glycerin.  To  preserve  the  preparation  permanently, 
proceed  as  in  (6).  To  do  away  with  the  macerating  action  of  the 
ammonium  picrate,  add  2  per  cent  of  a  1  per  cent  osmic-acid  solu- 
tion to  the  fixing  bath. 

60.  Methyl  Green. — This  is  one  of  the  best  of  the  nuclear  anilin 
stains.     It  is  particularly  valuable  because  it  often  instantly  stains 
the  chromatin  of  nuclei  in  fresh  tissues.     Use  in  strong  aqueous 
solutions,  acidulated  to  about  1  per  cent  with  acetic  acid.     It  does 
not  give  a  satisfactory  chromatin  stain  if  the  tissue  has  been  fixed 
in  acetic  acid  or  mixtures  containing  it.     It  follows  pure  corrosive- 
sublimate  solution  admirably. 

61.  Methyl  Violet. — This  stain  is  commonly  used  in  0.5  to  2  per 
cent  aqueous  solutions  for  staining  bacteria,  nuclei,  and  amyloid.     It 
may  often  be  substituted  for  gentian  violet. 

62.  Muci-Carmine  (Mayer). — 

Carmine 1.0  gram 

Aluminium  chloride 0.5  gram 

Distilled  water 2     c.c. 

Alcohol,  50  per  cent 100      c.c. 

Mix  in  the  order  given;  heat  gently  till  the  fluid  darkens  (about 
2  minutes) ;  filter  after  24  hours.  To  use,  dilute  with  5  to  10  volumes 
of  water.  The  stain  (3  to  10  minutes)  is  specific  for  mucus-containing 
cells. 

63.  Muci-Hematein  (Mayer).— 

Hematein 0.2  gram 

Glycerin 40      c.c. 

Aluminium  chloride 0.1  gram 

Distilled  water .  .  .  60  .    c.c. 


232  Animal  Micrology 

Rub  up  the  hematein  in  a  mortar  with  the  glycerin  and  the  alu- 
minium chloride,  then  add  the  water.  It  stains  in  from  3  to  10 
minutes,  mucin  appearing  blue.  If  a  drop  or  two  of  nitric  acid  is 
added,  its  nuclear  staining  capacity  is  enhanced. 

64.  Neutral  Red. — Neutral  red  is  used  widely  as  an  intra-vitam 
stain.     It  is  a  good  stain  for  cytoplasmic  granules,  and  in  some 
cases  for  mucus-cells.     For  intra-vitam  staining  it  may  be  used  in  the 
same  way  as  methylen  blue  (with  the  omission  of  fixation).     For 
staining  fixed  material,  a  1  per  cent  or  stronger  aqueous  solution  is 
employed.     Granules  are  stained  orange  red   (bright  red  in  acid 
medium,  yellow  in  alkaline  medium).     Rosin  finds  that  in  nerve 
cells  stained  in  neutral  red  (followed  by  water,  acid-free  alcohols, 
xylol,  and  balsam)  nucleoli  and  NissPs  granules  are  stained  red,  the 
rest  of  the  cell  yellow. 

65.  Orange  G. — This  is  an  excellent  cytoplasmic  stain  and  is 
often  used  on  sections  as  a  contrast  to  carmine,  hematoxylin,  and 
safranin.     It  is  especially  good  as  a  counterstain  in  tissues  of  verte- 
brate embryos.     Griibler's  orange  G  is  the  most  reliable.     It  should 
be  used  in  saturated  aqueous  solution.     The  solution  does  not  keep 
very  well. 

66.  Orcein  (Unna's  method  for  elastic  fibers) . — •• 

Orcein  (Griibler's) 1  gram 

Hydrochloric  acid 1  c.c. 

Absolute  alcohol 100  c.c. 

Sections  are  stained  in  a  watch-glass  or  porcelain  dish.  The 
dish  is  warmed  over  a  flame  or  in  an  oven  until  the  stain  becomes 
thick  through  the  evaporation  of  the  alcohol.  Rinse  the  stained 
sections  thoroughly  in  70  per  cent  alcohol,  wash  in  water,  run  up 
through  the  alcohols,  clear  in  xylol,  and  mount  in  balsam.  Elastic 
fibers  should  appear  dark  brown,  connective  tissue  a  pale  brown. 
Nuclei  may  be  brought  out  by  staining  in  Unna's  polychrome 
methylen-blue  solution  after  washing  the  sections  in  water. 

67.  Paracarmine  (Mayer's).— 

Carminic  acid 1.0  gram 

Aluminium  chloride 0.5  gram 

Calcium  chloride < 4.0  grams 

Alcohol,  70  per  cent 100     c.c. 


Some  Standard  Reagents  and  Their  Uses  233 

Paracarmine  is  an  excellent  stain  for  large  objects.  It  does  not 
overstain  ordinarily.  The  stained  tissue  is  washed  in  70  per  cent 
alcohol.  In  case  overstating  occurs  add  2 . 5  per  cent  glacial  acetic 
acid  or  0.5  per  cent  aluminium  chloride  to  the  alcohol  used  for 
washing.  Objects  to  be  stained  should  not  have  an  alkaline  reaction 
nor  contain  limy  materials. 

68.  Picric  Acid. — Picric  acid  is  widely  used  as  a  contrast  stain 
with  carmine,  hematoxylin,  etc.     It  is  best  manipulated  as  a  stain 
by  adding  a  little  to  each  of  the  alcohols  used  in  dehydrating,  after 
application  of  the  nuclear  stain.     However,  if  acid  alcohol  is  to  be 
used,  the  picric  acid  should  be  used  only  in  the  grades  above  the 
acid  alcohol.     It  may  be  employed  in  staining  entire  objects  as  well 
as  sections.     See  also  remarks  on  washing  under  22. 

69.  Picro-Carmine. — 

Ammonium  hydrate 5  c.c. 

Distilled  water 50  c.c. 

Carmine 1  gram 

When  dissolved  add  picric  acid  (saturated 

aqueous  solution) 50  c.c. 

Expose  to  air  and  light  for  two  days,  then  filter.  A  few  crystals 
of  picric  acid  should  be  added  to  the  alcohols  used  for  dehydration 
after  staining. 

Picro-Fuchsin. — See  47. 

Pyridine-Silver  Method  (Ranson's  Cajal). — See  p.  75. 

70.  Pyrogallol. — Tissues  which  have  been  fixed  in  Hermann's 
or  in  Flemming's  fluid  for  24  to  36  hours  may  be  treated  (without 
previous  washing)  with  a  weak  solution  of  pyrogallol  or  with  crude 
pyroligneous  acid.     Lee  (MicrotomisV s  Vade-Mecum)  recommends 
the  pyrogallol  as  much  preferable.     Tissues  should  remain  in  the 
fluid  from  1  to  24  hours,  depending  upon  size.    The  result  is  a 
black  stain  which  colors  both  nucleus  and  cytoplasm.     If  desired, 
an  additional  chromatin  stain   may  be  employed.     Safranin  (72) 
for  24  hours  is  recommended;   decolorize  slightly  with  very  dilute 
acid    alcohol.     The    stain   is   excellent   for   cytological    work    (for 
"sphere,"  etc.). 


234  Animal  Micrology 

71.  Resorcin-Fuchsin  (Weigert's  elastic  tissue  stain). — • 

Basic  fuchsin,  1  per  cent  aqueous  solution.  .....   100  c.c. 

Resorcin,  2  per  cent  aqueous  solution 100  c.c. 

Heat  the  mixture  in  a  porcelain  dish  and  while  boiling  add  25  c.c. 
of  liquor  ferri  sesquichlorati;  stir  and  keep  boiling  for  2  to  5  minutes. 
Cool  and  filter.  Throw  away  the  liquid.  Dry  the  precipitate 
which  remains  on  the  filter-paper  thoroughly  in  a  porcelain  dish 
over  a  water  or  sand  bath.  Return  the  dried  precipitate  together 
with  the  filter-paper  to  the  first  porcelain  dish,  add  200  c.c.  of  95  per 
cent  alcohol,  and  boil.  Remov'e  the  paper  when  the  precipitate  is  dis- 
solved off.  Cool,  filter,  and  replace  the  alcohol  lost  through  evapora- 
tion, up  to  200  c.c.  Add  4  c.c.  of  hydrochloric  acid.  The  stain 
works  best  after  formalin-fixed  material. 

Stain  the  section  20  minutes  to  an  hour  in  this  solution.  Wash 
in  alcohol,  dehydrate  in  absolute  alcohol,  clear,  and  mount  in  balsam. 
Elastic  fibers  should  appear  dark  blue  on  a  clear  background.  If 
desired,  the  sections  may  also  be  stained,  either  before  or  after  the 
staining  of  the  elastic  fibers,  with  one  of  the  carmine  or  hematoxylin 
stains. 

72.  Safranin. — Safranin  is  one  of  the  most  important  of  the  basic 
anilin  dyes.     Read  carefully  the  remarks  on  anilin  stains  under  30. 

Safranin 1  gram 

Anilin  water  (see  30) 90  c.c. 

Alcohol,  95  per  cent 10  c.c. 

Filter  before  using.  Griibler's  "Safranin  O"  is  the  most  reliable 
dye.  Sections  of  tissue  fixed  in  Hermann's  or  Flemming's  solution 
are  left  in  the  stains  for  from  24  to  48  hours.  Decolorize  as  directed 
under  30. 

73.  Safranin  and  Gentian  Violet. — This  is  a  combination  that 
is  almost  indispensable  in  the  study  of  cell  problems,  especially  sper- 
matogenesis.     For  formulae  of  stains  see  48  and  72.     Tissues  are 
best  fixed  in  Flemming's  or  Hermann's  solutions.     Stain  thin  sections 
for  36  to  48  hours  in  the  safranin;  differentiate  in  alcohol  very  slightly 
acidulated  (see  30),  then  stain  for  5  to  10  minutes  in  the  gentian  solu- 
tion and  transfer  the  sections  to  Gram's  solution  (see  under  48)  for 


Some  Standard  Reagents  and  Their  Uses  235 

1  to  3  hours.  Finally  differentiate  in  absolute  alcohol.  As  soon  as 
purple  clouds  have  ceased  to  come  from  the  sections  in  absolute 
alcohol,  they  should  be  transferred  to  clove  oil  for  a  few  minutes  and 
thence  to  xylol.  The  clove  oil  seems  to  intensify  the  safranin  in  the 
chromatic  granules,  but  too  prolonged  an  immersion  in  clove  oil 
extracts  the  gentian  violet. 

74.  Scharlach  R. — A  saturated  solution  of  the  dye  in  equal  parts 
of  70  per  cent  alcohol  and  acetone  is  used.     This  is  a  specific  stain 
for  fat.     For  example,  cover-slip  preparations  are  fixed  in  formalin 
vapor  for  5  or  10  minutes,  stained  5  minutes  in  the  Scharlach  R  solu- 
tion, rinsed  in  70  per  cent  alcohol,  washed  in  water,  counterstained 
with  alum-hematoxylin  or  methylen  blue,  washed  in  water,  and 
mounted  in  glycerin-jelly.     Frozen  sections  of  formalin-fixed  material 
may  be  treated  in  much  the  same  way. 

With  any  evaporation  of  the  alcohol  a  precipitate  forms,  hence 
staining  should  be  done  in  a  tightly  closed  vessel.  After  staining 
with  alum-hematoxylin,  if  the  sections  are  put  into  a  1  per  cent 
aqueous  solution  of  acetic  acid  for  3  minutes,  the  contrast  is  sharper. 

75.  Silver  Nitrate. — The  nitrate-of-silver  method  is  used  largely 
as  an  impregnation  method  for  work  on  nerve  tissue  and  for  demon- 
strating intercellular  substances  and  outlining  boundaries  of  cells  in 
the  epithelial  coverings  of  membranes,  etc.     Wash  the  fresh  tissue 
in  distilled  water,  then  place  it  for  2  to  5  minutes  in  0. 5  to  1  per  cent 
aqueous  solution  of  silver  nitrate.     Rinse  in  distilled  water,  then 
expose  the  tissue  to  bright  sunlight  in  water  or  glycerin  (or  in  70  per 
cent  alcohol,  if  it  be  mounted  in  balsam)  until  a  brown  coloration 
appears.     Temporary  mounts  should  be  made  in  glycerin.     For 
applications  see  pp.  71-74. 

76.  Sudan  IH. — This  is  a  specific  stain  for  fat.     See  remarks  on 
p.  147.     A  saturated  alcoholic  solution  is  used  (5  to  10  minutes). 
Wash  rapidly  in  alcohol.     Since  alcohol  is  a  solvent  of  fat,  too  long  an 
immersion   will   destroy  the   preparation.     Mount   in   glycerin   or 
glycerin-jelly.     The  tissue  should  have  been  fixed  previously  in 
Miiller's  fluid  (9)  or  other  medium  which  does  not  dissolve  fat. 

77.  Thionin. — This  is  an  excellent  stain  for  chromosomes  when 
used  in  saturated  aqueous  solution  for  about  5  minutes.     After 


236  Animal  Micrology 

corrosive-sublimate  fixation  it  is,  when  used  dilute  for  10  to  15 
minutes,  a  specific  stain  for  mucin  (mucin  red,  everything  else  blue). 
Van  Giesen's  Stain.— See  47. 

78.  Weigert-Pal  Stain  for  Medullated  Nerve  Fibers. — 

Solution  I: 

Hematoxylin 1  gram 

Absolute  alcohol 10  c.c. 

Distilled  water. 90  c.c. 

Lithium  carbonate  (1  part  of  a  saturated 

aqueous  solution  to  80  of  distilled  water)  1  c.c. 

Solution  II: 

Potassium  permanganate 0 . 25  gram 

DistiUed  water 100        c.c. 

Solution  III;   mix  just  prior  to  using: 

Oxalic  acid,  1  per  cent  aqueous  solution  ...     50       c.c. 
Potassium  sulphate,    1   per   cent   aqueous 

solution 50        c.c. 

Tissues  should  have  been  fixed  in  Miiller's  fluid  or  in  10  per  cent 
formalin,  hardened  in  alcohol,  and  sectioned.  Stain  sections  until 
black  (6  to  24  hours)  in  solution  I.  Wash  well  in  water  to  which  a 
few  drops  of  lithium  carbonate  have  been  added.  Transfer  to 
solution  II  and  leave  until  the  gray  matter  of  the  nervous  tissue 
becomes  brown  (J  to  2  minutes).  Rinse  in  water  and  decolorize  in 
solution  III  until  the  gray  matter  of  the  tissue  becomes  a  light 
brown  and  the  white  matter  a  steel  blue  (\  to  1  minute).  Each 
section  must  be  carefully  watched  to  get  a  satisfactory  result. 
Wash  in  running  water  or  in  several  changes  of  water,  dehydrate, 
clear,  and  mount  in  balsam.  If  desired,  a  counterstain  of  alum- 
cochineal  may  be  given  before  the  final  dehydration. 

79.  Wright's  Stain  (for  blood  and  for  the  malarial  parasite). — 
See  memoranda  5  and  6,  pp.  110-111. 

in.     NORMAL  OR  INDIFFERENT  FLUIDS 
(For  Fresh  Tissues) 

80.  Aqueous  Humor. — Obtained  by  puncturing  the  cornea  of  a 
freshly  excised  beef's  eye.     A  small  amount  may  readily  be  obtained 
by  means  of  a  capillary  pipette  from  the  eye  of  a  freshly  killed  frog. 


Some  Standard  Reagents  and  Their  Uses  237 

Amniotic  fluid  from  pig  or  cow  fetuses  is  a  serviceable  fluid  for  the 
examination  of  fresh  tissues. 

81.  Blood  Serum. — Blood  is  allowed  to  clot  and  after  24  hours 
the  serum  is  poured  off.     If  necessary  it  may  be  further  freed  of 
blood  cells  by  means  of  a  centrifuge.     The  serum  will  keep  for  only  a 
day  or  two.     Schultze's  iodized  serum  made  by  saturating  blood  serum 
with  iodine  is  sometimes  classed  as  an  indifferent  fluid,  but  it  is  really 
a  dissociating  fluid. 

82.  Locke's  Solution.— See  p.  137. 

83.  Normal  Saline. — 

Sodium  chloride 0.7  to  0.9  gram 

Distilled  water 100  c.c. 

84.  Ringer's  Solution. — 

Sodium  chloride 0. 80  part 

Calcium  chloride  (anhydrous) 0.02  part 

Potassium  chloride 0.02  part 

Sodium  bicarbonate 0.02  part 

Distilled  water 100  parts 

Dextrose  (may  be  left  out) 0 . 10  part 

The  following  formula  is  probably  better  adjusted  to  tissues  of 
warm-blooded  animals: 

Sodium  chloride 0 . 900  part 

Calcium  chloride  (anhydrous) 0 . 024  part 

Potassium  chloride 0 . 042  part 

Potassium  bicarbonate 0 . 020  part 

Distilled  water 100         parts 

Ringer's  solution  corresponds  more  nearly  to  normal  blood  serum 
than  does  normal  saline  and  is  therefore  less  likely  to  produce  dis- 
tortions in  tissue  elements. 

IV.    DISSOCIATING  FLUIDS 

85.  Bichromate  of  Potassium. — A  0 . 2  per  cent  aqueous  solution 
is  commonly  used.     Nerve  cells  of  the  spinal  cord  and  also  various 
epithelia  dissociate  wrell  in  it  (2  to  3  days). 

86.  Caustic  Potash. — A  solution  of  35  parts  in  100  parts  of  water 
is  often  used  for  isolating  fibers  of  smooth  muscle  or  heart  fibers. 


238  Animal  Micrology 

It  acts  by  rapidly  destroying  the  connective  tissue  (20  to  30 minutes). 
Examination  of  the  tissue  is  made  by  mounting  it  in  the  dissociating 
fluid.  If  water  is  added,  the  tissue  will  be  destroyed.  Usually  only 
temporary  preparations  are  made  in  this  fluid,  but  tissues  may  be 
made  permanent  by  neutralizing  the  alkali  by  means  of  acetic  acid. 
Digestion  Method.— See  pp.  79,  268. 

87.  Gage's  Formaldehyde  Dissociator. — 

Formalin 0.5  c.c. 

Normal  saline  solution 250      c.c. 

Good  for  epithelia  and  for  nerve  cells. 

88.  Hertwig's  Macerating  Fluid.— See  p.  78. 

89.  MacCallum's  Macerating  Fluid. — • 

Nitric  acid 1  part 

Glycerin 2  parts 

Water 2  parts 

This  fluid  is  recommended  for  heart  muscle  of  adults  or  embryos. 
Hearts  should  remain  in  it  from  8  hours  to  3  days,  according  to  size. 
The  method  is  valuable  for  showing  the  arrangement  of  cardiac 
muscle  fibers. 

90.  Ranvier's  One-Third  Alcohol. — This  is  one  of  the  common- 
est as  well  as  one  of  the  best  macerating  fluids.     It  is  simply  a  30 
per  cent  alcohol.     Epithelia  will  macerate  in  it  sufficiently  in  24  hours. 
A  still  weaker  alcohol  (20  to  25  per  cent)  is  used  for  isolating  the 
nerve  fibers  of  the  retina. 

91.  Sodium  Chloride. — A  10  per  cent  solution  of  sodium  chloride 
is  excellent  for  tendon,  etc.     It  dissolves  the  cement  substance  of 
epithelial  cells  and  of  connective  tissue.     As  a  stain,  a  saturated 
aqueous  solution  of  picric  acid  (stain  for  24  to  36  hours)  followed, 
after  thorough  washing  in  water,  by  a  dilute  alcoholic  solution  of 
acid  fuchsin  gives  excellent  results. 

V.  DECALCIFYING  FLUIDS 

Tissues  are  fixed  in  Zenker's  or  other  fluid,  thoroughly  washed,  and 
hardened  for  at  least  24  hours  in  alcohol  before  decalcification. 

92.  Chromic  Acid. — Chromic  acid  diluted  to  1  per  cent  or  in  com- 
bination with   other  fluids  is  frequently  used  for  decalcification. 


Some  Standard  Reagents  and  Their  Uses  239 

Chromic  acid,  1  gram;  water,  200  c.c.;  nitric  acid,  2  c.c.,  is  a  mixture 
widely  used.  It  decalcifies  well  but  acts  more  slowly  than  the  10 
per  cent  nitric-acid  mixture.  Bone  should  first  be  hardened  in 
Miiller'sfluid(9). 

93.  Nitric  Acid. — A  10  per  cent  solution  of  metric  acid  in  70 
per  cent  alcohol  may  be  used.    If  nitric  acid  is  used  for  young  or  fetal 
bones,  it  is  advisable  to  use  only  1  part  of  the  acid  to  99  parts  of  the 
alcohol.    After  washing  out  in  70  per  cent  alcohol,  the  decalcified  bone 
may  be  kept  in  95  per  cent  alcohol. 

94.  Phloroglucin  Method. — This  is  a  rapid  method.    Young 
bones  may  be  decalcified  in  half  an  hour  and  old  and  hard  ones  in  a 
few  hours.    Teeth  require  a  somewhat  longer  time.     Phloroglucin 
itself  does  not  decalcify,  but  protects  the  tissue  from  the  action  of  the 
strong  nitric  acid.    One  gram  of  phloroglucin  is  dissolved  in  10  c.c. 
of  pure  non-fuming  nitric  acid  with  the  aid  of  gentle  heat.    Ten  c.c. 
of  nitric  acid  in  100  c.c.  of  water  is  added  to  the  mixture.    Wash 
thoroughly  and  stain  in  Delafield's  hematoxylin.     After  staining 
leave  sections  in  tap  water  for  12  hours. 

95.  Picric  Acid. — A  solution  kept  fully  saturated  is  useful  for 
delicate  bones.     It  stains  and  decalcifies  the  tissue  at  the  same  time. 
Wash  in  70  per  cent  alcohol. 

96.  Von  Ebner's  Fluid.— 

Alcohol,  95  per  cent 500      c.c. 

Water 100      c.c. 

Sodium  chloride 2.5  grams 

Hydrochloric  acid 5.5  c.c. 

This  is  an  excellent  fluid  for  bone  because  in  it  the  ground  sub- 
stance of  the  bone  does  not  swell  up.  Sections  are  best  examined 
in  a  10  per  cent  solution  of  sodium  chloride. 


240 


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APPENDIX  D 

PREPARATION  OF  MICROSCOPICAL  MATERIAL  FOR  A 
GENERAL  COURSE  IN  ZOOLOGY 

(In  addition  to  the  methods  enumerated  here,  see  also  II,  chap,  x,  and 
chap,  xiii.) 

PROTOZOA 

a)  Cultures. — Amebae,  etc.,  may  usually  be  obtained  in  quanti- 
ties sufficient  for  class  use  by  the  following  method  recommended 
by  H.  S.  Jennings  (Journal  of  Applied  Microscopy,  VI,  No.  7,  p.  2406). 

A  number  of  glass  dishes  measuring  8  or  9  inches  in  diameter  by 
3  inches  deep  are  crowded  full  of  water  plants  (especially  Cerato- 
phyllum  and  Elodea),  filled  with  water,  and  the  plants  allowed  to 
decay.  Keep  the  dishes  in  warm,  light  places.  In  two  or  three 
weeks  the  layers  of  plants  at  the  surface  of  the  water  will  be  covered 
with  a  brown  slime  which  should  be  examined  occasionally  under 
the  microscope  for  the  desired  forms.  The  scum  that  appears  on  the 
surface  of  the  water  consists  mainly  of  bacteria  upon  which  amebae 
largely  feed.  They  will  be  found  most  frequently  in  the  slime  that 
immediately  surrounds  the  plant  tissue.  Since  they  frequently  last 
only  two  or  three  days  in  a  culture,  to  insure  material  for  class  work, 
a  number  of  cultures  must  be  made  at  different  dates  and  from 
different  localities.  Other  protozoa  such  as  Arcella,  Difflugia, 
Carchesium,  Stentor,  etc.,  will  also  be  found  in  the  cultures. 

As  soon  as  the  amebae  appear  in  such  cultures,  several  days 
before  they  are  desired  for  use,  Smith  (American  Naturalist,  XXXIX 
[1905],  467)  skims  off  the  brown  scum  and  puts  it  in  small  bacteria 
dishes  (4X1J  inches)  with  enough  water  to  fill  the  dish  about  1  inch 
deep.  He  adds  a  little  of  the  decaying  vegetable  matter  from  near 
the  surface  of  the  original  culture,  covers  the  dishes  and  keeps  them 
in  a  warm  place  but  out  of  direct  sunlight.  In  this  way  numerous 
large  active  specimens  may  often  be  obtained. 

259 


260  Animal  Micrology 

Pieces  of  frog  or  mussel  allowed  to  decay  for  about  10  days  in  pond 
water  will  usually  afford  an  abundance  of  a  small  species  of  ameba. 

Barker  keeps  amebae  and  paramecia  from  dying  out  by  adding 
a  sheet  of  fish  food  whenever  the  culture  begins  to  be  depleted. 
For  a  pure  culture  method  see  Kofoid,  Transactions  of  the  American 
Microscopical  Society,  XXXIV,  No.  4  (October,  1915). 

Chilomonas  and  Infusoria  usually  appear  in  a  few  days  in  cultures 
of  hornwort  and  partly  decayed  water-lily  leaves  packed  in  bacteria 
dishes  as  for  amebae,  but  with  proportionately  more  water. 

Paramecium  may  be  kept  from  dying  out  by  keeping  bits  of 
stale  bread  in  cultures.  A  culture  of  pond  water  and  bread  will 
usually  develop  large  numbers  of  paramecia  in  from  a  week  to  ten 
days.  See  also  last  paragraph  under  Euglena. 

Euglena  will  be  found  in  some  of  the  cultures,  but  usually  not 
in  any  quantities  before  the  end  of  four  or  five  weeks.  They  appear 
along  the  side  of  the  dish  toward  the  light.  Stephenson  finds  that  a 
few  grams  of  pulverized  rice  covered  with  pond  water  provides 
an  abundance  of  Euglena  in  from  ten  days  to  two  weeks. 

Turner  boils  20  grams  of  dry  quince  seed  for  half  an  hour  in  Ij 
liters  of  distilled  water,  then  passes  the  thick  exudate  which  is  given 
off,  together  with  the  water,  through  a  wire  sieve  (Eimer  and 
Amend,  No.  80).  He  then  makes  up  the  volume  to  2|  liters  with 
distilled  water,  sterilizes  it  and  places  it  in  a  stoppered  bottle. 
The  medium  will  keep  for  months.  Cultures  made  by  inoculating 
tubes  or  flasks  of  the  medium  with  Euglena  will  keep  for  a  year, 
and  specimens  can  be  obtained  in  considerable  numbers  at  any 
time  after  four  weeks.  Cultures  will  keep  longer  in  a  thicker 
medium  but  will  not  reproduce  rapidly. 

Standard  masses  for  use  in  experimental  work  may  be  prepared 
by  evaporating  the  exudate  to  dryness  and  making  up  solutions 
with  distilled  water.  A  0.2  per  cent  solution  of  the  dried  exudate 
seems  to  furnish  the  optimum  density.  Mold  frequently  invades 
the  cultures,  but  will  not  grow  in  a  density  of  0 . 2  per  cent  or  less. 
Mold  growths  a*re  less  likely  to  occur  in  cultures  which  have  been 
rendered  slightly  alkaline.  Cultures  should  be  kept  at  room  tempera- 
ture and  in  a  moderately  lighted  place. 


Preparation  of  Microscopical  Material  261 

Paramecia  and  other  infusoria  will  live  and  reproduce  for  about 
two  months  in  the  medium,  feeding  upon  Euglena  and  bacteria. 

Carchesium  and  Vorticella  are  frequently  found  on  decaying 
duckweed  (Lemna)  and  hornwort  (Ceratophyllum).  To  secure  a 
culture,  have  a  more  plentiful  supply  of  water  than  for  amebae. 
Professor  Walton  tells  me  that  he  always  finds  a  supply  of  Epistylis 
on  the  shells  of  fresh-water  snails. 

Didinium,  a  form  which  feeds  largely  on  paramecia,  is  highly 
recommended  by  Mast  (Science,  December  20,  1912,  p.  871)  as 
of  great  value  in  biological  study.  It  is  easy  to  obtain  (in  para- 
mecia cultures),  shows  the  phenomena  of  fission  and  encystment 
with  particular  clearness,  and  has  a  remarkable  method  of  feeding. 
Didinia  can  be  kept  indefinitely  in  the  encysted  state,  and  when 
wanted  for  study  will  appear  within  24  hours  to  a  few  days  in  active 
form  if  introduced  into  a  vigorous  culture  of  paramecia. 

Opalina  may  be  obtained  readily  by  killing  a  frog  with  chloro- 
form and  slitting  open  the  large  intestine.  Examine  scrapings  of 
the  epithelial  wall  in  normal  saline  (reagent  83,  Appendix  B). 

Sporozoa. — Gregarina  may  be  found  in  the  alimentary  canal  of 
the  meal  worm  or  the  cockroach  and  Monocystis  in  the  male  repro- 
ductive organs  of  the  earthworm.  They  are  best  studied*  in  normal 
saline.  If  it  is  desired  to  stain  and  mount  specimens,  they  may  be 
fixed  in  corrosive-acetic  (reagent  15,  Appendix  B)  for  5  minutes, 
washed  thoroughly  in  35  per  cent  alcohol,  to  which  a  little  tincture  of 
iodine  has  been  added,  and  stained  with  Ehrlich's  triple  stain  (reagent 
42),  or  hematoxylin  and  acid  fuchsin  (reagents  52  and  45). 

Herpetomonas  may  be  obtained  from  the  intestine  of  the  fly 
and  of  the  squash  bug,  and  Trypanosoma  from  the  blood  of  the  rat. 

Volvox. — Volvox  globator,  the  form  commonly  described  in 
textbooks,  is  found  in  the  early  spring,  often  in  great  abundance, 
in  small  permanent  pools  which  contain  duckweed  and  Riccia.  A 
smaller  and  less  desirable  form,  Volvox  aureus,  may  be  found  later 
in  the  spring  and  throughout  the  summer  in  the  same  pools.  When 
water  from  such  pools,  together  with  a  small  amount  of  the  water 
plants,  is  placed  in  bacteria  dishes,  so  arranged  that  one  side  is 
strongly  exposed  to  light,  the  volvox  present  will  collect  after  a  few 


262  Animal  Micrology 

hours  at  the  edge  of  the  water  on  the  lighted  side  of  the  dish.  If 
the  contents  of  the*  vessels  which  contain  volvox  are  kept  in  as 
near  the  natural  condition  of  the  pond  as  possible,  the  organisms 
may  be  kept  alive  for  some  weeks  in  the  laboratory.  Tap  water  is 
injurious  to  them.  Avoid  having  too  much  decaying  material  in  the 
water,  although  some  is  essential.  Keep  in  glass-covered  dishes 
near  windows  (out  of  direct  sunlight)  in  as  cool  a  place  as  possible. 
Any  considerable  rise  in  temperature  beyond  that  of  the  original 
pond  will  result  in  their  death.  Small  Crustacea  feed  upon  volvox 
and  will,  if  present  in  any  considerable  numbers,  soon  exterminate 
them.  The  sexual  stages  are  more  likely  to  be  found  in  the  ooze 
at  the  bottom  of  cultures. 

Because  of  the  uncertainty  of  obtaining  living  volvox  at  any 
stated  time,  it  is  well  to  have  an  abundance  of  the  material  preserved 
in  5  per  cent  formalin.  Such  preserved  specimens  show  the  flagella 
more  distinctly  than  do  living  ones.  See  Smith,  The  American 
Naturalist,  XLI,  No.  481  (1907),  p.  31. 

6)  Quieting  infusoria. — 1.  Let  sufficient  water  evaporate  from 
under  the  cover  to  permit  the  latter  to  press  lightly  upon  the  animals. 
Guard  against  too  great  evaporation  of  water  or  the  infusoria  will 
be  crushed 

2.  Entanglement  in  fibers  of  cotton,   etc.,   sometimes  proves 
efficacious. 

3.  A  small  amount  of  gelatin,  or,  better,  cherry-tree  gum,  dis- 
solved in  water  makes  a  viscous  mass  which  is  often  useful  in 
retarding  their  motions.     A  bit  of  white  of  egg  may  be  used  in  the 
same  way. 

4.  Animals  may  be  narcotized  by  means  of  a  small  drop  of  very 
dilute  alcohol  (preferably  methyl  alcohol)  or  chloretone  (about  one 
drop  of  a  1  per  cent  solution  to  10  drops  of  water).     (Chloretone  is 
manufactured  by  Parke,  Davis  &  t2o.,  of  Detroit,  Mich.     For  its 
use  as  an  anaesthetic  in  biological  work  see  Journal  of  Applied 
Microscopy,  V,  2051.) 

c)  Feeding. — -Place  finely  pulverized  carmine  or  indigo  under 
the  cover-glass.  The  colored  powder  rapidly  accumulates  in  the 
food  vacuoles.  In  such  a  preparation  the  action  of  the  cilia  of 


Preparation  of  Microscopical  Material  263 

infusoria  is  also  indicated  by  the  rapid  movement  of  the  particles 
in  the  vicinity  of  the  animal.     See  also  memorandum  4,  p.  110. 

d)  Staining. — For  intra-vitam  staining  see  reagents  59a,  32,  and 
64,  Appendix  B. 

To  see  cilia  of  infusoria  treat  the  animal  with  very  dilute  iodine 
solution  or  a  drop  of  a  dilute  solution  of  tannin. 

To  see  the  macronucleus  and  the  micronucleus  use  a  drop  of  a  2  per 
cent  solution  of  acetic  acid,  or,  better,  methyl  green  (reagent  60, 
Appendix  B). 

e)  Permanent  mounted  preparations. — Benedict's  method  is  as 
follows: 

"Smear  a  glass  slide  with  albumen  fixative,  as  in  preparing  for  the 
mounting  of  paraffin  sections.  Then  place  on  the  surface  of  the  film 
of  fixative  a  drop  or  two  of  water  containing  the  form  which  it  is 
desired  to  stain.  Let  nearly  all  the  water  evaporate  by  exposure  to 
the  air  of  the  room  until  only  the  film  of  fixative  remains  moist. 
The  slide  can  now  be  immersed  in  Gilson  or  any  other  fixing  reagent, 
and  then  passed  through  the  alcohols,  stains,  etc.,  in  the  same  way 
that  mounted  sections  are  handled. 

"I  have  had  no  difficulty  in  getting  preparations  of  paramecium 
by  this  method,  with  very  little  distortion  of  the  body  and  any 
kind  of  staining  desired.  By  this  method  students  can  prepare 
in  ten  minutes  very  satisfactory  preparations  of  protozoa  for 
demonstration  of  nuclei,  etc." — Journal  of  Applied  Microscopy, 
VI,  2647. 

For  fixation  of  protozoa  Calkins  (Journal  of  Experimental 
Zoology,  I,  No.  3,  1904)  uses  saturated  aqueous  solution  of  corrosive 
sublimate  to  which  10  per  cent  of  glacial  acetic  acid  is  added.  See 
also  reagent  20  (formol  sublimate),  p.  214.  Barker  fixes,  washes, 
stains,  destains,  dehydrates,  clears,  and  mixes  protozoa  with  balsam, 
all  in  homeopathic  vials.  After  each  operation  material  is  allowed 
to  settle  well.  Reagents  are  pipetted  off. 

Plankton,  in  general,  are  well  fixed  if  passed  directly  into  fresh 
Zenker's  fluid  to  which  a  few  drops  of  1  per  cent  solution  of  osmic 
acid  have  been  added.  Under  such  treatment  ciliates  remain 
expanded. 


264  Animal  Micrology 

SPONGES 

To  isolate  the  spicules  of  calcareous  sponges,  boil  a  bit  of  the 
sponge  in  5  per  cent  solution  of  caustic  potash  for  a  few  minutes. 

Fairly  thick  transverse,  longitudinal,  and  tangential  sections  of 
Grantia  showing  spicules  in  the  tissues  are  useful.  Make  these  with 
an  old  razor  or  sharp  scalpel.  To  hold  the  object  while  sectioning, 
place  it  between  two  pieces  of  pith  or  cork.  For  a  careful  study  of 
the  relations  of  the  two  systems  of  canals  in  the  body  wall,  thinner 
sections  are  necessary.  To  prepare  these  it  is  best  to  decalcify  (2  per 
cent  chromic  acid,  24  to  36  hours)  the  sponge  and  cut  celloidin  or 
paraffin  sections  on  the  microtome,  although  fairly  good  sections  may 
be  made  by  hand.  They  should  be  dehydrated  and  mounted  in 
balsam  if  permanent  preparations  are  desired;  if  not,  they  may  be 
examined  in  glycerin. 

To  color  the  collar  cells  use  an  aqueous  solution  of  anilin  blue. 

Spicules  of  siliceous  sponges  are  isolated  by  treating  bits  of  the 
sponge  with  strong  nitric  acid  or  a  mixture  of  nitric  and  hydrochloric 

acid. 

COELENTERATES 

Hydra  should  be  sought  for  in  spring-fed  pools.  In  the  autumn 
they  are  found  most  frequently  on  smooth  dead  leaves  which  are 
completely  submerged.  Material  should  be  collected  and  placed 
in  battery  jars  or  larger  glass  jars,  which  are  then  filled  with  fresh, 
clear  water,  and  placed  in  a  fairly  light  place,  but  not  too  near  a 
window.  Put  a  small  amount  of  hornwort  or  Chara  in  each  jar. 
In  a  few  hours  (12  to  36)  the  hydra  will  be  found  attached  to  the  sides 
of  the  vessel  and  to  the  plants.  They  may  readily  be  kept  in  the 
laboratory  throughout  the  winter  if  glass  plates  are  placed  over  the 
jars  to  prevent  excessive  evaporation  and  the  temperature  is  not 
allowed  to  go  below  freezing.  Fresh  water  should  be  added  from  time 
to  time  to  make  up  for  evaporation.  In  case  their  supply  of  food 
(Cyclops,  Daphnia,  and  other  small  Crustacea)  is  exhausted  it  should 
be  renewed  by  skimming  out  from  other  aquaria  the  small  forms 
upon  which  the  animal  feeds  and  putting  them  in  the  hydra  jars. 

Keeping  hydra  in  the  dark  at  somewhat  lower  temperature  for 
several  days  favors  the  formation  of  spermaries  and  ovaries. 


Preparation  of  Microscopical  Material  265 

For  staining  and  mounting  entire  see  p.  95.  Kill  in  the  same  way 
for  sectioning.  The  most  instructive  sections  are  (1)  transverse 
sections,  (2)  longitudinal  sections  through  the  mouth  and  a  bud,  and 
(3)  sections  showing  the  sexual  organs.  Stain  in  bulk  with  hema- 
toxylin  (reagent  52,  Appendix  B),  imbed  in  paraffin,  using  the  method 
for  delicate  objects  (p.  53),  and  after  the  paraffin  has  been  removed 
from  the  sections,  stain  them  for  a  few  seconds  in  acid  fuchsin. 
Dehydrate  and  mount  in  the  usual  way. 

The  sections  are  much  more  satisfactory  if  the  hydra  have  been 
placed  in  small  stender  dishes  filled  with  filtered"  water  (not  dis- 
tilled) and  kept  from  food  for  a  week  or  ten  days  before  killing.  This 
eliminates  the  metabolic  products  and  oil  globules  which  ordinarily 
obscure  the  details  of  structure. 

To  Stain  the  Nematocysts  of  Living  Hydra,  place  several  of  the 
animals  in  a  small  stender  dish  of  water  which  has  been  tinted  a 
sky  blue  through  the  addition  of  methylen-blue  solution  made  up 
as  follows: 

Methylen  blue 1.0  gram 

Castile  soap 0.5  gram 

Water 300     c.c. 

After  two  hours  the  hydra  may  be  transferred  to  fresh  water;  the 
nematocyst  cells  are  stained  a  deep  blue.  (Method  of  Little,  Journal 
of  Applied  Microscopy,  VI,  2216.) 

To  Discharge  Nematocysts  drum  on  the  cover-glass  gently  with  a 
pencil.  By  using  a  very  small  opening  to  the  diaphragm  they  are 
usually  sufficiently  distinct  without  staining. 

For  Other  Polypoid  Forms  the  methods  given  for  hydra  will 
answer  in  most  cases. 

For  Collecting  Free-Swimming  Medusoid  Forms  full  directions 
will  be  found  in  Brook's  Invertebrate  Zoology. 

Compound  Hydrozoa  should  be  placed  alive  into  the  cells  which 
they  are  to  occupy  when  mounted;  1  per  cent  formic  acid  is  then 
added  drop  by  drop  to  the  sea  water.  After  the  animals  have  been 
killed,  the  fluid  is  replaced  by  glycerin- jelly  and  the  cover-glass  is  put 
in  place.  Another  method  is  to  kill  the  animals  slowly  by  adding  a 


266  Animal  Micrology 

few  crystals  of  chloral  hydrate  from  time  to  time  to  the  small 
vessel  of  sea  water  containing  them. 

Small  Jellyfish  may  be  fixed  and  hardened  in  1  per  cent  osmic 
acid  and,  stained  or  unstained,  mounted  in  cells. 

Anemones,  Medusae,  and  other  delicate  marine  forms  may 
usually  be  killed  in  the  expanded  condition  by  means  of  magnesium 
sulphate.  Success  lies  in  securing  a  quick  diffusion  of  a  quantity  of 
the  sulphate  through  the  water  without  causing  mechanical  disturb- 
ance of  the  animal  to  be  anaesthetized.  Griffin  accomplishes  this  by 
tying  a  considerable  quantity  of  the  magnesium  sulphate  in  a  piece 
of  cheesecloth  and  hanging  it  over  the  dish  of  sea  water  containing 
the  animals  in  such  a  way  that  the  bottom  of  the  bag  barely  dips  into 
the  water.  Mayer's  method  of  anaesthetizing  medusae  by  carbon 
dioxide  is  also  often  applicable  to  other  sensitive  contractile  forms. 

PLANARIA 

Look  for  planarians  on  the  under  sides  of  stones  in  small  streams 
of  running  water.  They  are  usually  examined  alive.  To  see  them 
thrust  out  the  proboscis,  keep  them  from  food  for  a  few  days  and 
then  feed  them  on  dead  flies.  Planaria  which  have  been  kept  in  the 
laboratory  for  months  display  the  internal  organs  much  more  clearly 
than  freshly  captured  ones. 

If  it  is  desired  to  study  stained  specimens,  for  preparation  see  p.  97. 

To  Kill  Planaria  with  Pharynx  Protruded,  Cole  (Journal  of 
Applied  Microscopy,  VI,  2125)  recommends  covering  them  in  a  watch- 
glass  with  a  1  per  cent  aqueous  solution  of  chloretone  until  they  are 
immobilized  and  then  rapidly  transferring  them  to  5  per  cent  formalin. 
Other  fixing  agents  than  formalin  can  be  used. 

TREMATODES 

The  most  easily  obtained  forms  are  those  found  in  the  lungs, 
intestine,  or  bladder  of  frogs.  A  good  form  for  study  is  occasionally 
found  in  the  liver  of  the  cat.  Search  for  it  in  the  bile  passages. 
Fix  trematodes  in  corrosive  sublimate,  wash  out  with  alcohol  to 
which  tincture  of  iodine  has  been  added,,  and  stain  for  24  hours  in 
alum-cochineal  (reagent  28,  Appendix  B)  or  carmalum  (reagent  35). 


Preparation  of  Microscopical  Material  267 

As  with  planaria,  they  should  be  compressed  between  two  glass 
slides  (see  p.  97).  To  kill  trematodes  in  a  distended  condition, 
Barker  flattens  out  each  individual  on  a  glass  slide  or  in  a  watch- 
glass  with  a  camel's  hair  brush  and  floods  it  with  the  killing  agent. 

If  the  large  liver  fluke  of  the  sheep  (Fasciola  hepatica)  can  be 
obtained,  both  the  alimentary  canal  and  the  excretory  system  may 
be  injected  with  India  ink  or  with  finely  powdered  carmine  in 
water.  For  injection  a  very  fine-pointed  cannula  with  rubber  cap 
is  used,  or  the  manipulator  may  operate  the  cannula  by  simply 
blowing  through  it.  The  excretory  system  is  injected  through  an 
incision  made  with  a  sharp-pointed  scalpel  in  the  median  line  near 
the  hinder  end  of  the  animal.  For  the  alimentary  canal  the  incision 
should  be  made  about  1  mm.  to  one  side  of  the  median  line.  When 
the  injection  is  completed,  flatten  the  animal  somewhat  between  two 
slides  (see  p.  97),  harden  in  95  per  cent  alcohol  for  12  to  24  hours,  then 
dehydrate,  clear,  and  mount  in  balsam. 

Larval  Stages  may  frequently  be  found  in  the  so-called  "liver" 
of  pond  snails. 

CESTODES 

Near  large  cities  an  unlimited  supply  of  the  sheep  tapeworm 
(Monieza)  can  usually  be  secured  from  slaughter  houses.  Ample 
supplies  can  ordinarily  be  obtained  from  dogs,  or,  less  frequently,  from 
cats.  Tapeworms  can  be  kept  alive  for  a  considerable  length  of  time 
in  tepid  water.  The  most  instructive  portions  to  mount  are  scolex, 
sexually  mature,  and  terminal  proglottids.  For  fixing  and  staining 
use  the  same  methods  as  for  distomes.  La  Rue  finds  that  carmine 
stains  are  better  for  trematodes  and  hematoxylins  for  cestodes. 
The  scolex  should  not  be  compressed.  To  kill  cestodes  in  an  extended 
condition,  Barker  wraps  the  living  worm  around  a  glass  slide,  than 
immerses  the  slide  in  the  killing  reagent.  The  worm  is  removed  as 
soon  as  killed. 

To  Find  Cysticerci,  open  the  body  cavity  of  a  rabbit  and  look  for 
large  whitish  bodies  imbedded  in  the  peritoneum  or  liver  (the  cysti- 
cercus  of  T.  serrata).  Likewise,  the  cysticercus  of  T.  crassicollis 
may  be  found  in  the  liver  of  the  mouse.  If  a  cysticercus  is  found, 
its  outer  wall  should  be  slit  open  in  order  to  show  the  reversed  scolex. 


268 


Animal  Micrology 


NEMATODES 

See  memorandum  20,  p.  136.     Nematodes  occur  frequently  in 
the  intestines  of  pigs,  dogs,  cats,  and  rabbits. 

Trichinella. — The  simplest  way  to  obtain  it  is  to  apply  for  infected 
pork  to  the  government  inspector  whose  headquarters  are  to  be 

found  near  all  large 
slaughter  houses  in  cities. 
Bits  of  the  infected  muscle 
should  be  teased  and 
flattened  out  in  a  com- 
pressor (Figs.  73  and  74) 
until  a  favorable  area  has 
been  found .  The  flattened 
tissue  may  then  be  dehydrated  and  mounted  unstained  or  it  may  be 
stained  in  hematoxylin  (reagent  52,  Appendix  B) .  Better  results  will 
be  obtained  if  the  material  is  fixed  for  from  4  to  6  hours  in  Carnoy's 
fluid  (reagent  2)  before  dehydrating  or  staining.  If  desired,  the 
tissue  may  be  sectioned  in  celloidin  or  paraffin. 


PIG.  73. — Compressor 


FIQ.  74. — Compressor  Used  by  the  Government  Bureaus  for  Meat  Inspection 

To  Demonstrate  Living  Trichinellae,  Barnes  (American  Monthly 
Microscopical  Journal,  XIV,  104)  subjects  small  bits  of  trichinized 
muscle  to  a  mixture  of  3  grains  of  pepsin,  2  drams  of  water,  and  2 
minims  of  hydrochloric  acid  for  about  three  hours  at  body  tempera- 
ture with  occasional  shaking.  When  the  flesh  and  cysts  are  dis- 
solved, the  liquid  is  poured  into  a  narrow  glass  vessel  and  allowed  to 


Preparation  of  Microscopical  Material  269 

settle.     The  live  trichinae  may  be  withdrawn  with  a  pipette  from 
the  bottom  of  the  fluid  and  examined  on  a  warm  stage. 

ROTIFERS 

Rotifers  will  usually  be  found  in  abundance  in  some  of  the  labora- 
tory aquaria  on  the  lighted  side  of  the  vessel.  For  ordinary  class 
work  they  are  best  studied  alive.  They  are  difficult  to  preserve 
properly.  Full  directions  for  killing  and  preserving  will  be  found 
in  Jenning's  paper,  "Rotatoria  of  the  United  States,"  U.S.  Fish 
Commission  Bulletin  (1902),  p.  277. 

To  Quiet  Rotifers,  Cole  (Journal  of  Applied  Microscopy,  VI, 
2179)  anaesthetizes  them  by  adding  from  time  to  time  a  drop  of  1  per 
cent  aqueous  solution  of  chloretone  to  the  water  on  the  slide  in  which 
the  animals  are  being  examined. 

BRYOZOA 

They  may  be  treated  in  the  same  way  as  compound  hydrozoa. 
Plumatella  may  frequently  be  found  in  shallow  fresh-water  streams 
on  the  under  side  of  flat  rocks;  Pectinatella}  in  rivers  and  streams 
on  the  upper  surface  of  mussel  shells,  etc. 

STARFISH 

Barker's  technique  for  Pedicellaria  is  as  follows :  Boil  the  aboral 
part  of  a  ray  from  a  f ormalin-preserved  starfish  in  5  per  cent  caustic 
soda  for  from  3  to  5  minutes.  Wash  quickly  and  thoroughly  in 
water,  stain  in  water-soluble  eosin,  wash  in  acid  (acetic)  alcohol, 
dehydrate,  and  mount. 

EARTHWORMS 

Earthworms  are  best  collected  on  warm,  rainy  nights  when  they 
may  be  found  extended  on  the  surface  of  the  ground  near  their  bur- 
rows. They  are  most  plentiful  in  old  gardens  or  rich  lawns.  A 
lantern  and  a  pail  are  the  only  implements  necessary.  Earthworms 
may  frequently  be  found,  however,  in  large  numbers  on  the  surface 
of  the  ground  on  cloudy  days  immediately  after  prolonged  hard  rain. 
In  winter  living  ones  can  nearly  always  be  found  under  manure 
piles. 


270  Animal  Micrology 

To  Prepare  Earthworms  for  Class  Work,  secure  good-sized 
specimens,  wash  them  in  water,  and  place  them  in  a  vessel  containing 
moist  filter-paper.  Put  only  a  few  worms  in  each  dish  and  adjust  the 
cover  so  as  to  admit  a  little  air.  After  12  to  24  hours  it  is  well  to 
remove  any  dead  or  injured  specimens  and  to  change  the  filter-paper. 
The  dish  should  be  kept  from  direct  sunlight  in  a  cool  place.  After 
two  or  three  days  the  grit  and  dirt  in  the  alimentary  canal  will  have 
been  passed  out  and  its  place  taken  by  paper  which  the  worms  have 
eaten.  They  are  then  ready  to  kill  and  preserve  or  section. 

Place  the  worms  in  a  flat  vessel  and  pour  on  sufficient  water  to 
cover  them.  During  the  next  two  hours  add  a  little  alcohol  from  time 
to  time  until  the  strength  of  the  liquid  is  increased  to  about  8  or  10 
per  cent.  Then  wash  all  mucus  from  the  body  of  the  worms  and 
replace  them  in  10  per  cent  alcohol  until  they  no  longer  respond  to 
pricking  or  pinching  with  forceps.  Transfer  them  to  50  per  cent 
alcohol  for  several  hours,  keeping  them  straightened  out  as  much  as 
possible;  then  to  70  per  cent  alcohol  for  12  hours,  followed  by  95  per 
cent  alcohol  for  24  hours.  Preserve  finally  in  70  per  cent  alcohol. 

Chromic-Acid  Method.- — Although  requiring  considerable  more 
work  in  preparation,  specimens  hardened  in  chromic  acid  are  so  much 
superior  to  alcoholic  ones  for  general  dissection  purposes  that  the 
extra  trouble  is  well  worth  while.  The  worms  are  anaesthetized  as 
in  the  preceding  method,  but  from  10  per  cent  alcohol  they  are 
injected  with  1  per  cent  aqueous  solution  of  chromic  acid  and  then 
immersed  in  it  for  4  hours.  While  working  with  the  chromic  acid  the 
hands  and  wrists  should  be  coated  with  vaseline. 

Keeping  the  worms  extended  and  submerged  in  1  per  cent 
chromic  acid  in  a  large  shallow  dish,  inject  the  acid  into  the  body 
cavity  slowly,  about  half  an  inch  behind  the  clitellum,  and  again 
near  the  posterior  end  of  the  body.  Avoid  piercing  the  alimentary 
canal.  The  injection  is  not  complete  until  the  worm  is  turgid  along 
its  entire  length.  The  worms  must  be  kept  straight  and  untwisted 
while  in  the  chromic  acid.  Remove  them  at  the  end  of  4  hours  (a 
longer  time  in  the  acid  will  make  them  brittle)  and  wash  thoroughly 
in  running  water  until  the  yellow  color  is  gone  (12  to  16  hours). 
Remove  them  to  50  per  cent  alcohol  for  2  days,  then  to  70  per 


Preparation  of  Microscopical  Material  271 

cent  alcohol  for  2  or  3  days,  and  finally  preserve  in  fresh  70  per  cent 
alcohol. 

For  injection  a  water-pressure  apparatus  (Fig.  35)  is  best.  The 
reservoir  A  should  be  placed  about  4  feet  above  the  compression 
chamber  B.  The  cannula  should  be  made  of  a  piece  of  quarter-inch 
glass  tubing  with  one  end  drawn  out  to  a  very  fine  bore  and  so  broken 
as  to  leave  a  sharp  point  and  edge  for  piercing  the  body  wall  of  the 
worm. 

For  sectioning,  the  preliminary  steps  are  the  same  as  in  the 
alcohol  method,  but  from  10  per  cent  alcohol  the  worms  should  be 
placed  into  Zenker's  fluid  (reagent  6,  Appendix  B)  for  4  to  6  hours. 
For  washing,  etc.,  follow  the  directions  given  in  the  discussion  of  the 
reagent.  To  facilitate  penetration  of  the  fluid,  it  is  well  to  slit  open 
the  body  cavity  of  the  worm  in  places  that  are  not  to  be  sectioned. 
The  most  instructive  sections  are  cross-sections  of  the  middle  of 
the  body  and  sagittal  sections  of  the  anterior  end  which  include 
the  pharynx.  The  worms  may  be  stained  in  bulk  (24  to  36  hours) 
in  borax-carmine  (reagent  33)  or  hematoxylin  (reagent  52)  before 
sectioning. 

Entire  nephridia.  together  with  a  small  part  of  the  septum  which 
they  traverse  should  be  carefully  dissected  out,  stained  in  borax- 
carmine  (reagent  33),  dehydrated,  cleared,  and  mounted  in  balsam. 

An  ovary  should  be  removed  entire,  stained  with  borax-carmine, 
dehydrated,  cleared,  and  mounted  in  balsam. 

A  testis  should  be  treated  in  the  same  way  as  an  ovary.  Tease 
it  in  the  balsam  before  adding  the  cover-glass. 

To  Keep  Earthworms  Alive  in  Winter,  Jennings  (Journal  of 
Applied  Microscopy,  VI,  2412)  places  them,  immediately  after 
collection,  into  bacteria  dishes  (9  in.  in  diameter  by  3  in.  deep) 
between  folds  of  muslin  which  is  kept  damp  but  not  dripping  wet. 
Not  more  than  a  dozen  worms  should  be  placed  in  one  dish  and  the 
cloth  should  be  changed  or  washed  at  least  every  two  weeks.  The 
worms  may  be  fed  on  leaves,  etc.,  from  time  to  time. 

To  Immobilize  Earthworms  for  study  of  circulation  of  the  blood 
under  the  microscope  or  projection  lantern,  Cole  (Journal  of  Applied 
Microscopy,  VI,  2125)  places  them  in  a  0. 2  per  cent  aqueous  solution 


272  Animal  Micrology 

of  chloretone  for  3  or  4  minutes.     Such  worms  may  be  slightly 
compressed  between  two  slides. 

To  Examine  Corpuscles  of  the  Coelomic  Fluid,  expose  the  worms 
for  a  minute  or  two  to  the  vapor  of  chloroform.  The  coelomic 
fluid  exudes  through  dorsal  pores.  Touch  a  cover-glass  to  the 
fluid  and  mount. 

The  Setae  Can  Be  Isolated  by  boiling  a  bit  of  the  tissue  containing 
them  in  a  solution  of  caustic  potash.  When  isolated,  dry  them  and 
mount  in  balsam. 

LEECH 

Leeches  are  obtained  from  fresh-water  pools,  streams,  and  marshes, 
but  to  get  sufficient  numbers  for  class  use  it  is  usually  necessary  to 
purchase  them  from  dealers.  Live  leeches  intended  for  dissection 
may  be  killed  with  chloroform.  Cross-sections  prepared  in  the 
same  way  as  for  earthworms  are  very  instructive. 

ARTHROPODS 

For  Mounting  Small  Crustacea  see  III,  A,  chap.  xiii. 

To  Quiet  Small  Crustacea  for  Microscopical  Examination  (Cole, 
Journal  of  Applied  Microscopy,  VI,  2180)  place  them  in  a  watch- 
glass  containing  2  parts  of  1  per  cent  chloretone  and  5  parts  of  water. 
The  same  treatment  is  useful  for  the  larvae  of  insects.  Some,  such  as 
the  nymph  of  the  dragon  fly,  will  require  more  chloretone. 

For  Various  Dissections  and  Parts  of  Insects  see  II,  chap.  x. 

For  Mounting  Insects  Entire  (beetles,  mosquitoes,  gnats,  aphids, 
larvae,  etc.)  as  microscopic  preparations,  and  for  mounting  muscle, 
wings,  heads,  legs,  scales,  antennae,  etc.,  see  chap.  xiii. 

Live  nymphs  of  the  dragon  fly  are  especially  valuable  for  study 
under  the  compound  microscope  because  they  show  very  clearly 
the  valvular  action  of  the  heart,  the  tracheal  gills  and  tracheae,  and 
the  brain  and  its  relation  to  the  eyes.  The  heart  is  located  well 
toward  the  posterior  end  of  the  abdomen  between  the  main  tracheal 
trunks.  Cole  (Journal  of  Applied  Microscopy,  VI,  2274)  recom- 
mends that  the  animals  be  anaesthetized  by  subjecting  them  to  a 
1  per  cent  aqueous  solution  of  chloretone.  , 


Preparation  of  Microscopical  Material  273 

MOLLUSKS 

Gills  of  the  Fresh-Water  Mussel  may  be  fixed  in  corrosive 
sublimate  (reagent  14,  Appendix  B)  for  from  20  to  30  minutes,  washed 
out  in  water  and  then  hi  dilute  alcohol  to  which  tincture  of  iodine  has 
been  added.  Make  cross-sections  hi  paraffin,  stain  hi  dilute  hema- 
toxylin  (reagent  52),  and  mount  in  the  ordinary  way. 

Cross-Sections  of  the  Entire  Mussel  are  valuable  to  show  the 
relations  of  the  gills,  kidneys,  and  heart.  Wedge  the  valves  apart 
slightly  and  immerse  the  animal  for  24  hours  hi  1  per  cent  chromic 
acid  (reagent  11).  Wash  out  thoroughly  hi  running  water  and 
transfer  the  specimens  to  70  per  cent  alcohol  for  two  or  three  days  or 
until  needed.  To  section,  remove  both  valves,  place  the  animal  on  a 
board,  and  with  a  razor  cut  transverse  sections.  These  are  to  be 
examined  with  the  naked  eye  or  with  a  dissecting  lens. 

To  Kill  Snails  in  an  Expanded  Condition,  put  them  into  a  vessel 
of  cold  water,  then  run  a  layer  of  hot  water  on  to  the  surface  of 
the  cold  water.  See  that  the  vessel  is  full  of  water  and  cover  it 
with  a  glass  plate  to  exclude  the  air. 

For  Lingual  Ribbon  of  the  Snail  see  memorandum  7,  p.  100. 

AMPfflOXUS 

Specimens  must  ordinarily  be  secured  from  dealers.  The  animals 
should  be  stained  entire  hi  borax-carmine  (reagent  33,  Appendix  B) 
and  sectioned  hi  celloidin.  The  most  instructive  sections  are  cross- 
sections  of  a  female  with  well-developed  gonads  and  longitudinal 
sections  of  small  individuals.  Mounts  of  entire  small  specimens 
should  also  be  made. 

VERTEBRATA 

For  any  of  the  tissues  of  vertebrates  which  teachers  may  desire 
to  prepare,  ample  directions  are  given  in  Appendix  C. 

For  Demonstration  of  Circulation  of  the  Blood  in  the  frog,  see 
chap.  xiv. 


APPENDIX  E 

TABLE  OF  EQUIVALENT  WEIGHTS  AND  MEASURES 
WEIGHTS,  METRIC  AND  AVOIRDUPOIS 

1  kilo  =  1,000  grams  =  1  liter  of  water  at  its  maximum  density  =  2.2  pounds.    « 
1  gram  =  l  cubic  centimeter  of  water  at  its  maximum  density  =15. 43 

grains  =  0 . 035  ounce. 
1  pound  =  453 . 59  grams. 
1  ounce  =  28. 35  grams. 
1  grain  (Troy)  =  0.065  gram. 
1  dram  =  1.77  grams. 

WEIGHTS,  METRIC  AND  APOTHECARY'S 

1  kilo  =  1,000  grams. 

1  gram  =  15 . 43  grains  =  0 . 032  ounce. 

1  pound = 373 . 24  grams. 

1  ounce  =  31 . 10  grams. 

1  dram  =  3 . 89  grams. 

1  scruple  =  1 . 30  grams. 

1  grain  =  0.065  gram. 

MEASURES  OF  LENGTH,  METRIC  AND  ENGLISH 

1  meter  =  1,000  millimeters  =  39 .37  inches. 

1  centimeter  =  0.394  inch. 

1  millimeter  =  0 . 039  inch. 

1  yard = 0 . 9 14  meter. 

1  foot  =  30. 48  centimeters. 

1  inch  =  2 . 54  centimeters  =  25 . 40  millimeters. 

LIQUID  MEASURES,  METRIC  AND  APOTHECARY'S 

1  liter  =1,000  cubic  centimeters  =  2. 11  pints. 
1  cubic  centimeter =0.034  fluid  ounce  =16. 23  minims. 
1  gallon  =  128  ounces  =  3 . 79  liters.        » 
1  pint  =  16  ounces  =  473. 18  cubic  centimeters. 
1  fluid  ounce  =  8  fluid  drams  =  29. 57  cubic  centimeters. 
1  fluid  dram  =  60  minims  =  3. 70  cubic  centimeters. 

274 


Equivalent  Weights  and  Measures  275 

THERMOMETERS 

To  reduce  degrees  Fahrenheit  to  degrees  Centigrade  use  the  formula, 
C  =  5/9(F— 32).  For  example,  if  the  number  of  degrees  Fahrenheit  is  77, 
then  C  =  5/9  (77—32)  =  25  degrees.  Or,  for  instance,  to  reduce  —31  degrees 
Fahrenheit  to  Centigrade,  C  =  5/9  (-31-32)  =  5/9X  -63=  -35  degrees. 

To  reduce  degrees  of  Centigrade  to  degrees  of  Fahrenheit  use  the  formula 
F=9/5C+32.  For  example,  if  the  number  of  degrees  Centigrade  is  25, 
then  F  =  (9/5  X  25) +32  =  77  degrees.  Or,  to  reduce  -  35  degrees  Centigrade 
to  Fahrenheit,  F=  (9/5 X  -35)+32  =  -31  degrees. 


APPENDIX  F 
REFERENCES 

Only  a  very  limited  bibliography  is  given.  A  full  one  will  be 
found  in  Gage's  The  Microscope.  Above  all,  for  desirable  special 
methods  the  student  is  advised  to  look  through  the  articles  in  cur- 
rent journals  which  cover  the  field  of  his  own  researches. 

Of  American  periodicals  the  Anatomical  Record  and  the  Trans- 
actions of  the  American  Microscopical  Society  make  a  special  point 
of  technique.  In  England  the  Journal  of  the  Royal  Microscopical 
Society  does  the  same.  Other  periodicals  which  give  prominence 
to  technique  are:  the  Zeitschrift  fur  wissenschaftliche  Mikroskopie 
und  fur  mikroskopische  Technik;  the  Zoologischen  Anzeiger;  Anato- 
mischer  Anzeiger,  and  the  Biologisches  Centralblatt. 

The  following  books  should  be  consulted  for  detailed  information 
on  points  of  zoological  and  histological  technique: 

Chamberlain,  Methods  in  Plant  Histology.     The  University  of  Chicago  Press, 

Chicago,  111.,  1915. 
Cole,  A  Manual  of  Biological  Projection  and  Anesthesia  of  Animals.     Neeves 

Stationery  Co.,  Chicago,  111.,  1907. 
Ehrlich,   Krause,   Moore,   Rosin,   and  Weigert,   Encyclopadie  der  mikro- 

skopischen  Technik.    Urban  und  Schwarzenberg,  .Berlin,  1910. 
Gage,  The  Microscope.    Cdmstock  Publishing  Co.,  Ithaca,  N.Y.,  1916. 
Hardesty,    Neurological    Technique.    The    University    of    Chicago    Press, 

Chicago,  111.,  1902. 

Lee,  The  Microtomist's  Vade-Mecum.    Blakiston,  Philadelphia,  1913. 
Mallory  and  Wright,  Pathological  Technique.     Saunders,  Philadelphia,  1915. 
Mann,  Physiological  Histology;    Methods  and  Theory.     Clarendon  Press, 

.Oxford,  1902. 

Wright,  Principles  of  Microscopy.     Constable,  London,  1907. 
Any  of  the  recent  textbooks  on  Histology. 


276 


INDEX 


INDEX 


Abbe,  190,  200. 

Abbot's  method  for  staining  spores  of 
bacteria,  118. 

Aberration:  chromatic,  183;  correction  of 
chromatic,  183;  correction  of  spherical, 
182;  spherical,  181. 

Absolute  alcohol:  preparing,  7;  testing 
for  water,  56. 

Accessory  chromosomes,  151. 

Acetic  acid,  17,  207;  for  contractile 
animals,  207. 

Acetic  alcohol,  208. 

Achromatic  objective,  187. 

Achromatism,  187. 

Acid  carmine,  146,  222. 

Acid  fuchsin,  224;  Altman's,  145;  methyl- 
green  method  for  mitochondria,  144; 
methyl-green  stain,  Auerbach's,  224; 
orange  G,  methyl-green  mixture,  Ehr- 
lich's,  223. 

Acid  magenta,  224. 

Acidophil  granules,  240. 

Adipose  tissue,  242. 

Affixing  sections,  23;  frozen,  70;  paraffin, 
41. 

Agminated  nodules,  246. 

Air-bubbles,  206. 

Albumen  fixative,  12,  23. 

Albuminized  water,  23. 

Alcohol,  7,  16,  17,  28,  208;  absolute,  7; 
absolute,  test  for  water  in,  55;  acetic- 
chloroform  mixture,  207,  and  corrosive 
sublimate,  208;  acid,  8,  24,  48;  alkaline, 
48;  and  ether,  8,  214;  ethyl,  13;  fixation, 
28 ;  gradesjof ,  7  ;  methyl,  13 ;  renewing, 
55;  wood,  13. 

Alcoholometer,  13. 

Algae,  mounting  medium  for,  100. 

Alimentary  canal,  245;  of  insects,  to 
mount,  79. 

Allen,  119,  120,  122,  139,  152,  153. 

Allen's  B-15  method,  149;  gelatin  method, 
122. 

Altman's  acid  fuchsin,  145. 

Alum-carmine,  218;   dahlia,  241. 

Alum-cochineal,  9,  218. 

Ambystoma,  122;  epidermal  cells  of,  142; 
mitosis  in  cells  of,  140,  142;  to  study 
living  cells  in,  143. 

Ameba,  259. 

Ammonia  copper  sulphate  solution,  195. 

Amphibia:  artificial  fecundation  in,  135; 
embryology  of,  119-23. 

Amphioxus,  273. 

Amphophil  granules,  240. 

Amyloid,  219,  246. 


Anatomical  material,  preservation  of,  31. 

Andrews,  125. 

Anemones,  266. 

Angular  aperture,  187. 

Anilin  blue,  orange  G,  and  acid  fuchsin, 

219;    dyes  avoided  in  celloidin  method, 

63;    oil,  22,  153;    stain,  acid,  basic,  20; 

stains,  20,  218;  stains,  decolorizing,  219; 

water,  219. 

Antennae,  to  mount,  99. 
Anthrax,  116,  118. 
Aorta,  241. 

Apathy's  celloidin  and  paraffin  method,  64. 
Apertometer,  187. 
Aphid,  to  kUl  and  mount,  98. 
Aplanatism,  187. 

Apochromatic  objective,  183,  188. 
Apparatus  required,  1. 
Aqueous  humor,  236. 
Arcella,  259. 

Archoplasm,  stains  for,  147. 
Areas  of  Cohnheim,  250. 
Areolar  tissue,  247. 
Artery,  241. 
Arthropods,  272. 
Ascaris,    polar    bodies,    fertilization    and 

cleavage  in,  136,  137. 
Assheton,  132. 
Auerbach's    fuchsin    methyl-green    stain, 

Aurantia,  20. 

Axial  illumination,  194. 

Axis  cylinder,  251. 

Axis  of  lens:    principal,  177;    secondary, 

177. 
Axone,  251. 

Bacillus,  115;  aerogenes  capsulatus,  118; 
of  anthrax,  116,  118;  of  bubonic  plague, 
118;  of  chancroid,  118;  coli  communis, 
118;  diphtheriae,  116,  118;  of  dysen- 
tery, 118;  of  glanders,  118;  of  influenza, 
118;  of  malignant  edema,  118;  mucosus 
capsulatus,  118;  proteus,  118;  pyano- 
cyaneus,  118;  of  tetanus,  118;  of  tuber- 
culosis, 118;  of  typhoid,  118. 

Bacteria:  cover-glass  preparations  of, 
112;  features  to  be  observed  in  study, 
ing,  115;  Gram's  method  of  staining, 
114;  hanging-drop  preparations  of, 
112,  115;  in  tissues,  112,  114;  material 
for  demonstrating,  115;  methylen  blue 
stain  for,  114;  mounting  from  fluid 
media,  112;  mounting  from  solid 
media,  113;  staining  and  mounting 
films,  112;  spores,  115,  118;  staining 
flagella,  118;  stains  for,  116. 


279 


280 


Animal  Micrology 


Bacterial  examination,  112. 

Balsam,  11,  22;  bottle,  4;  mounting  in, 
34,49;  mounts,  fading  of ,  58 ;  removing 
exuded,  57;  xylol-,  12. 

Bardeen,  156,  157;  freezing  microtome,  68. 

Barker,  101,  153,  263,  267. 

Basophil  granules,  240. 

Beetles,  to  mount  as  opaque  objects,  98. 

Beggiatoa,  115. 

Bell's  cement,  95. 

Benedict,  263. 

Bensley,  32,  143;  mitochondrial  methods, 
144,  145. 

Benzaldehyde  for  clearing  and  fixing 
whole  objects,  104. 

Benzol,  22. 

Benzopurpurin,  20. 

Bergamot  oil,  22,  153. 

Berlin  blue,  84. 

Bethe's  fluid,  229. 

Bibliography,  276. 

Bichloride  of  mercury,  208. 

Bichromate  of  potassium,  208;  dissociat- 
ing fluid,  237;  in  various  fixing  mixtures, 
209,  210. 

Binocular  loop,  Hardy,  189. 

Binocular  magnifier,  189. 

Binocular  microscope,  189. 

Bismarck  brown,  20,  146,  220. 

Black  pins  in  sections,  57. 

Bladder,  257. 

Blastoderm:  of  chick,  124;  of  fish,  139, 141. 

Bleaching,  24,  44. 

Bleu  de  Lyon,  20,  227. 

Blocks  for  mounting  celloidin,  63. 

Blood,  105-11;  and  blood-forming  organs, 
240;  clinical  examination  of,  107; 
corpuscles,  105;  corpuscles,  living,  105; 
cover-glass  preparation,  106,  240;  crys- 
tals, 105,  106;  currents,  to  observe, 
109;  dry  preparations  of,  106;  effects 
of  reagents  on,  105;  enumeration  of 
corpuscles,  107;  examination  of  fresh, 
105;  platelets,  105,  111;  rapid  method 
for,  107;  Schultze's  iodized  serum,  237; 
serum,  237;  serum,  Loeffler's,  117; 
to  study  in  sections,  110;  tests  for, 
106;  Wright's  stain  for,  110. 

Blotting-paper  method  of  reconstruction, 
157. 

Blue  color,  to  restore  in  injected  tissues,  90. 

Bone:  corpuscles,  242;  decalcifying,  81, 
243;  development  of,  243;  fibers  of 
Sharpey,  243;  grinding,  82;  Haversian 
canals  and  lamellae,  245;  isolation  of 
corpuscles,  245;  sectioning,  81;  stains 
for  developing,  220. 

Books  on  micro- technique,  276. 

Borax-carmine,  Grenacher's,  220. 

Borax-ferricyanide,  145. 

Bordeaux  red,  147,  221. 

Born,  154. 

Bouin's  picro-formol  fixing  fluid,  9,  29; 
fixing  with,  2$;  use  in  cyto'ogy,  139. 

Boveri,  30,  208. 


Brain  cells,  251;  sand,  251. 

Brittle    objects:     celloidin    and    paraffin 

method  for,  64;   paraffin-rubber  method 

for,  43. 
Bronchi,  256. 
Brooks,  265. 

Brownian  movement,  189. 
Bryozoa,  269. 
Burckhardt,  131. 
Burr,  131. 
Burrows,  138. 

Cajal's  method  for  neuro-fibrils,  75. 

Calcification,  test  for,  147. 

Calibration  of  microscope,  189. 

Calkins,  263. 

Calleja's  staining  fluid,  222. 

Camera  lucida,  189,  190;  Abbe,  190; 
Wollaston,  190. 

Cannulae,  glass,  90. 

Capillaries  and  small  vessels,  241. 

Capillaries,  Golgi  method  for,  73. 

Carbol-fuchsin  for  bacteria,  116. 

Carbolic  acid,  22,  32. 

Carbon  dioxide:  for  freezing,  67;  to 
estimate  minute  quantities  of,  153. 

Carborundum  points,  1. 

Carchesium,  259,  261. 

Cards,  record,  5. 

Carmalum,  Mayer's,  221. 

Carmine,  20;  Beale's,  221;  borax-,  220; 
muci-,  231;  picric  acid,  and  indigo- 
carmine,  222;  picro-,  233;  Schneider's 
acid,  222. 

Carnoy:  fixing  fluids,  103,  207;  Lebrun 
fixing  fluids,  139,  208. 

Carrel,  138. 

Cartilage,  243;  capsule  of,  243;  con- 
nective tissue  and  elastic  fibers  in,  243; 
elastic,  243;  glycogen  in,  243;  hyaline, 
243 ;  white  flbro-,  243. 

Cassia  oil,  22. 

Caustic  potash,  caustic  soda,  25. 

Cedarwood  oil,  22,  42. 

Celloidin,  12,  23;  blocks  for  mounting, 
63;  bottle,  4;  cement,  134;  hardening, 
62;  imbedding  a  number  of  objects  in, 
63;  removing  from  sections,  66;  sec- 
tions, affixing,  24;  sections,  to  transfer 
from  the  knife,  64;  serial  sections,  65. 

Celloidin  and  paraffin  methods  compared, 
63. 

Celloidin  method,  26,  59;  clearing  objects 
in  the  block,  64;  Gilson's  rapid,  66; 
in  cytology,  151;  length  of  time,  62. 

Celloidin-paraffin  method  for  brittle 
objects,  64. 

Cell:  dissection  of  living,  153;  walls,  to 
stain,  147. 

Cell-making,  93,  101 ;    deep,  101. 

Cells  of  Paneth,  245. 

Cellular  .structures,  tests  for  certain,  147. 

Celluloid:  for  corrosion  preparations.  91; 
plating  method,  65. 


Index 


281 


Centering  an  object  in  a  cell,  100. 

Centigrade  to  Fahrenheit,  to  reduce,  275. 

Central  illumination,  194. 

Central  nervous  system,  251. 

Centrosome,  140,  147. 

Cerebellar  cortex,  251. 

Cerebral  cortex,  251. 

Ceruminous  glands,  247. 

Cestodes,  267. 

Chamberlain,  276. 

Chancroid,  bacillus  of,  118. 

Chick:  embryology  of,   123-26;    marking 

anterior  end   of  young,    130;     window 

method  for,  138. 
Child,  122,  128. 
Chilomonas,  260. 
Chinese  black,  136. 
Chinolin  blue,  222. 
Chloretone,  262. 

Chloride  and  acetate  of  copper,  211. 
Chlorine,  Mayer's  bleaching  method,  44. 
Chloroform,  22;    in  celloidin  method,  61; 

in  paraffin  method,  53. 
Cholera,  spirillum  of,  118. 
Choroid,  249;  plexus,  252. 
Chrom-acetic^osmic     acid,     211;      acetic- 
formalin  mixture,  212. 
Chromatin,  tests  for,  147. 
Chromic  acid,  17,  211;  decalcifying  fluid, 

238;  washing  out,  17. 
Chromosomes,   stains   for,    147,    235;     to 

demonstrate  quickly,  151. 
Chrom-silver  method,  Golgi's,  71. 
Cicatricula,  124. 
Cinnamic  aldehyde,  22. 
Circulatory  system,  241. 
Cleaning  mixture,  57;    microscope  lenses, 

203,  205;    slides  and  covers,  56. 
learers,  21, 
Clearing,  21. 
Cleavage:    in  living  material,  136;    quick 

preparation  of  stages  in  teleosts,  136. 
Clinging  of  paraffin  sections  to  the  knife, 

Clitoris,  254. 

Clove  oil,  22;    in  minute  dissections,  80. 

Coccus,  115. 

Cochineal,  20;    alum,  9,  218;    and  Lyons 

blue,  50. 
Cochlea,    247;     nerve    fibers    and    nerve 

endings  of,  247. 
Coelenterates,  264,  265. 
Cole,  266,  269,  271,  272,  276. 
Collar  cells  of  sponges,  264. 
Collodion,  23;  instead  of  celloidin,  65. 
Colostrum,  257. 

Compound  microscope,    manipulation    of 

the,  203. 

Compressors,  268. 
Condenser,  191. 
Congo  glycerin,  80. 
Congo  red,  10,  52,  222. 


Conklin,  125,  129. 
Connective  tissues,  242-45. 
Contractile  animals  killing,  16,  207. 
Convex  or  converging  lens,  176. 
Cooler  for  paraffin  microtome,  47.~ 
Cooling  tissues,  153. 
Coplin  staining  jars,  4. 

Copper  chrome-hematoxylin   method   for 

mitochondria,  145. 
Coregonus,  141. 

Cornea,  249;  silver-nitrate  method  for,  74. 
Corneal  corpuscles  and  nerves,  249. 
Corneal  spaces  and  canaliculi,  249. 
Corpus  luteum,  254. 
Correction  collar,  191. 
Corrosion  of  injected  vessels,  25,  90,  91. 
Corrosive  sublimate,  8,  9,  17.  18,  20,  28, 

209,     212;      handling,    9;     in    various 

fixing   fluids,    208,    209,    213;     washing 

out,  18,  28. 
Cort,  100. 
Cover-glass,    1;    correction  for   thickness 

of,  191,  192;  standard,  191;  to  support, 

80. 

Cowdry,  143,  145. 

Cox  modification  of  the  Golgi  method,  73. 

Crayfish,  mitosis  in  testis  of,  140. 

Creosote,  22. 

Crescents  of  Gianuzzi,  245. 

Crooked  paraffin  ribbons,  45. 

Crown  glass,  183. 

Crumbling  of  objects  or  tissues,  43;    in 

paraffin,  45,  47. 

Crustacea:   small,  96;    to  quiet,  273. 
Crystal  violet,  144. 
Culture  slide,  115. 
Curare,  109. 

Cyanin,  222;    and  erythrosin  for  sperma- 
tozoa, 222. 

Cylindrical  end  bulbs,  252. 
Cysticerci,  to  find,  267. 
Cytological  methods,  139. 
Cytoplasmic  stains,  21. 

Damar,  gum,  22. 

Danchakoff's  mixture,  210. 

Danforth,  131. 

Daniel,  131. 

Dark-ground  illumination,  192. 

Day  lite  glass,  192. 

Dealcoholization,  22. 

Dealers,  4. 

Decalcification,  25;238,  fluids  for,  9,    239. 

Decolorization,  24. 

Dehydration,  18. 

Delicate  tissues,  42;    paraffin  method  for 
53. 

Demilunes  of  Heidenhain,  245. 
Demonstration  microscope,  192. 
Demonstration  ocular,  192. 
Desilicidation,  25. 


282 


Animal  Micrology 


Developing  bone,  to  stain,  220. 

Diapedesis,  109. 

Diaphragm,  192. 

Didinium,  261. 

Difflugia,  259. 

Digestive  organs,  245;  blood  vessels  of, 
245. 

Dilution,  rules  for,  7,  13. 

Diphtheria,  116,  118. 

Diplococci,  115. 

Diplococcus  intra  cellularis  meningitidis, 
118. 

Dipping  tube,  101. 

Discoidal  cleavage,  129. 

Dispersion,  182. 

Dissecting  instruments,  1. 

Dissection  of  living  cells,  153;  of  small 
embryos,  134. 

Dissections,  minute,  77. 

Dissociation,  15,  25;  by  means  of  formal- 
dehyde, 77;  fluids  for,  237;  general 
rules  for,  80. 

Dogiel's  methylen  blue  method,  229. 

Double  staining,  19,  50. 

Drawing,  159;  for  publication,  166;  in 
cytology,  166;  in  embryology,  165; 
in  histology,  166;  in  special  courses, 
164;  materials,  159;  methods,  159;  use 
of  camera  lucida  in,  168,  189;  use  of 
colored  crayons  in,  165;  table  for 
reconstruction  work,  157. 

Drawings:  arrangement  of,  163;  arrange- 
ment of,  for  reduction,  172;  combina- 
tion, 170;  flxing  pencil,  162;  for  half- 
tone reproduction,  170;  for  publication, 
167;  for  reproduction  in  color,  170,  171; 
ink,  161;  labeling,  164;  lettering  for 
publication,  164,  172;  pencil,  161; 
reduction  of,  168;  reproduction  of,  by 
line  process,  168;  reproduction  of 
pencil,  171;  reproduction  of  wash,  170; 
shading,  161,  166;  to  keep  clean,  163; 
wash,  162. 

Drop  method,  18,  152. 

Dropping  bottle,  58. 

Dry  areas  under  the  cover-glass,  57. 

Dry  mounts,  99. 

Ducts,  Golgi  method  for,  76. 

Duodenum  245. 

Duval,  130. 

Dysentery,  bacillus  of,  118. 


Ear:  external,  247;  middle,  247. 
Earthworm,    269;     alcohol    method    for, 

270;     chromic-acid    method    for,    270; 

keeping  alive  in  winter,  271;   nephridia 

of,  271;  ovary  of,  271;   sectioning,  271; 

testis    of,    271;     to    examine    coelomic 

corpuscles  of,  272;    to  immobilize,  271; 

to  isolate  setae  of,  272. 
Eau  de  Javelle,  25. 
Echinoderms,    artificial    fecundation    in, 

135. 
Eggs,    notes     on    Amphibian,     121;      of 

insects  in  dry  mounts,  99;    of  mouse, 

130-31. 


Ehrlich,  276;  -Biondi  triple  stain,  222; 
triple  stain,  223. 

Elastic  elements  of  blood  vessels,  241. 

Elastic  fibers,  244. 

Electrification  of  paraffin  sections,  47. 

Embalming,  31,  32.     ' 

Embryograph,  193. 

Embryological  methods,  119. 

Embryology:  amphibia,  119-21;  chick, 
123-26,  129;  frog,  119;  mouse,  130, 
131;  pig,  133;  rabbit,  131,  132;  rat, 
131,  133;  teleosts,  128,  129. 

Embryonic  shield,  128,  129. 

Embryonic  tissues,  cultivation  of,  removed, 
137. 

Embryos:  dissection  of  small,  134;  early 
stages  of  mammalian,  131;  human, 
134;  injection  of,  88,  89;  marking 
anterior  end  of  young  chick,  130; 
measuring  length  of,  128;  older  stages 
of  mammalian,  133 ;  whole  mounts  and 
sections  of  teleost,  128. 

Encircling  fibers,  244. 

Endothelial  cells,  248. 

Endothelium  of  blood  vessels,  241. 

Eosin,  10,  20,  50,  224. 

Eosinophil  granules,  240. 

Epidermis,  under-surface  view  of,  256. 

Epididymus,  254. 

Epistylis,  261. 

Epithelial  tissues,  248,  249. 

Epithelium:  ciliated,  248;  ciliated  colum- 
nar, 248;  columnar  and  glandular, 
248;  cubical,  248;  of  alimentary  tract, 
245;  pigmented,  248;  squamous  or 
pavement,  249;  stratified,  249;  tran- 
sitional, 249. 

Erlicki's  fluid,  210. 

Erythrocytes,  111. 

Erythrosin,  20,  224. 

Esophagus,  245. 

Ether:  and  alcohol,  8,  214;  for  freezing, 
69,  70. 

Euglena,  260. 

Euparal  as  a  mounting  and  preserving 
medium,  22,  54,  153. 

Eustachian  tube,  247. 

Evans,  89,  146. 

Evaporation  from  cells,  to  prevent,  101. 

Eycleshymer's  clearer,  63;  methods  of 
orienting  objects  in  celloidin,  127,  128. 

Eye,  249. 

Eyelid,  249. 

Eyepiece.     See  Ocular. 

Eye-Point,  184,  193. 

Fading  of  balsam  mounts,  58. 
Fahrenheit  to  Centigrade,  to  reduce,  275. 
Fallopian  tube,  254. 
Fat:  cells,  244;    intestinal  absorption  of, 

246;    tests  for,    147;    to  remove  from 

tissues,  148. 

Fecundation,  artificial,  135. 
Fenestrated  membrane,  244. 


Index 


283 


Fertilization  in  mammals,  130. 

Fetuses,  134. 

Fibrillar  connective  tissue,  244. 

Fibrin,  stained  preparation  of,  105. 

Fixation,  16,  30;   by  injection,  30. 

Fixing  and  hardening  agents,  207-18. 

Fixing:  general  rules  for,  27;  tissues  cut 
by  the  freezing  method,  70;  with  alco- 
hol, 28;  with  Bouin's  fluid,  29;  with 
formalin,  29;  with  Zenker's  fluid,  28. 

Flagella  of  bacteria,  staining,  118. 

Flatness  of  field,  193. 

Flatworms,  killing  and  mounting,  97,  100, 
266,  267. 

Flemming's  fixing  fluid,  211;  use  in 
cytology,  139. 

Flint,  91. 

Flint  glass,  183. 

Flukes,  100. 

Focal  point,  176. 

Foci,  conjugate,  175. 

Foliate  papillae,  247. 

Foot  and  Strobell,  method  of  photo- 
micrography, 150. 

Forceps,  1. 

Formalin,  8,  214;  fixing  with,  29;  for 
frozen  sections,  30;  in  various  fixing 
mixtures,  214,  215;  protecting  the 
skin  from,  32;  to  neutralize,  214; 
with  Zenker's  fluid,  209. 

Formic  glycerin,  132. 

Formol.     See  Formalin. 

Free  acid,  to  detect,  in  tissues,  148. 

Free-hand  sectioning,  33. 

Freezing  attachment  for  ether,  69. 

Freezing  method,  67;  for  fixed  tissues, 
70;  for  fresh  tissues,  69;  for  objects 
alcohol  injures,  70. 

Fresh  tissues,  26;  sectioning  by  the 
freezing  method,j59;  staining,  146. 

Friable  objects,  sectioning,  43,  44,  62. 

Frog  embryos:  fixing,  staining,  and 
mounting,  119-21;  section  method  for, 
119;  whole  mounts  of,- 122. 

Frozen  sections,  affixing,  70. 

Fuchsin:  acid,  20,  224;  and  methyl-green 
stain,  224;  and  picric  acid  stain,  224; 
basic,  224;  other  combinations  with, 
116,  144,  219,  222,  223,  224,  234. 

Gabbet's  method  for  tubercle  bacilli,  117. 
Gage,  26,  110,  147,  157,  216,  276;    congo- 

glycerin,  80;    formaldehyde  dissociator, 

238;   stain  for  glycogen,  148. 
Gall  bladder,  245. 
Gall  duct,  245. 
Ganglion  canaliculi,  252. 
Ganglion  cells,  252. 
Gastric  glands,  245. 
Gelatin  capsules  for  small  objects,  31. 
Gelatin  injection  mass,  83 ;   to  keep,  90. 
Gelatin  method,  Allen's,  122. 
Gelatinous  coats  of  eggs,  to  remove,  121. 
Gentian  violet,  20,  116,  225. 


Germinal  layers,  122. 
Germ-ring,  128,  129. 
Gilson's  fixing  fluid,  213;  use  in  cytology, 

139. 

Gilson's  rapid  celloidin  process,  66. 
Gizzard  of  cricket  or  katydid,  to  mount, 

79. 

Glanders,  116,  118. 
Glaser,  47. 

Glass,  marking,  1,  58. 
Glycerin,  21,  35;   mounting  in,  93,  94. 
Glycerin  jelly,  21,  96;   mounting  in,  96. 
Glycogen,  test  for,  148. 
Gnat,  to  kill  and  mount,  98. 
Goblet  cells,  245. 
Gold    chloride    225;     method    for    nerve 

endings,  76. 

Gold  size,  to  dilute,  94. 
Golgi  method,  71-73;    determining  which 

elements   will   be   impregnated   in,   73; 

permanent  preparations  in,  73. 
Gonococcus,  118. 
Graafian  follicle,  254. 
Gradual  change  of  liquids,   method   for, 

152. 

Graduated  cylinders,  4. 
Gram's  method,  114,  117,  118. 
Gram's  solution,  117,  225. 
Grandry's  corpuscles,  252. 
Grantia,  264. 
Graphs,  169. 

Grasshopper,  mitosis  in,  140. 
Grave,  47. 
Gregarina,  261. 
Gregory  88 

Grenacher's  borax-carmine,  220. 
Griffin,  266. 
Griggs,  58. 
Guild,  76. 
Guncotton,  23. 

Hair,  256;  development  of,  257;  elements 
of,  257;  follicle,  257;  renewal  of,  257. 

Half-tones:  from  drawings,  170,  171; 
from  photographs,  170. 

Hance,  40,  58. 

Hanging-drop  preparations,  115. 

Hard  objects,  81-82. 

Hardening,  16,  17. 

Harder's  glands,  249. 

Hardesty,  44,  73,  74,  172,  276. 

Harrison,  138. 

Head  of  fly,  to  mount,  98. 

Heart,  242;   valves  of,  242. 

Heart-beat  of  insect,  to  see,  272. 

Heidenhain's  iron-hematoxylin,  10,  51. 

Heliotype  method  of  reproducing  draw- 
ings, 171. 

Helly's  fluid,  209. 

Hematein,  20;   muci-,  231. 

Hematoidin  crystals,  106. 


284 


Animal  Micrology 


Hematoxylin,  20;  and  eosin,  double 
staining,  50,  62;  bluing  by  means  of 
tap-water  or  alkaline  alcohol,  55; 
Conkiin's  picro-,  225;  Delafleld's,  9, 
34,  48;  Ehrlich's  acid,  226;  Heiden- 
hain's  iron-,  10,  51,  226;  Mallory's 
phosphotungstic,  226;  precise  staining 
in  Delafleld's,  56;  ripening,  9,  20. 

Hemin,  crystals,  106. 

Hemocytometer,  108. 

Hemoglobin:  crystals  of,  105;  tests  for, 
148 

Hemolymph  gland,  242. 

Hepatic  interlobular  connective  tissue,  246. 

Hepatic  lobules,  246. 

Herbst  corpuscles,  252. 

Hermann's  fluid,  217. 

Herpetomonas,  261. 

Hertwig  macerating  fluid,  78. 

Heuser,  88,  134,  157,  158. 

His,  128,  156. 

Hollow  organs,  30. 

Holmgren's  canals,  153. 

Homogeneous  immersion  objective,  193. 

Hones,  43. 

Hoyer,  89. 

Hubbert,  58. 

Huber,  65,  76,  133,  155. 

Human  embryos,  134. 

Huygenian  ocular,  180. 

Hydra,  264;  killing  and  mounting,  95; 
sectioning,  265;  staining  and  mounting, 
265;  staining  nematocysts  of,  265; 
to  discharge  nematocysts  of,  265;  to 
secure  spermaries  and  ovaries  in,  264. 

Hydrofluoric  acid,  25. 

Hydrozoa,  265. 

Hypophysis,  252. 

Illumination,  194;    dark  ground,  192. 
Images:     defects   in,    181;     formation   of 

by  lenses,  177. 
Imbedding,  22;    in  celloidin,  59;    in  gum 

and  syrup,  67;   in  paraffin,  36. 
Impregnation   methods,   21. 
Incubation  of  hen's  egg,  123. 
Indifferent  fluids,  26,  236. 
Indulin,  240. 

Indulinophil  granules,  240. 
Infiltration  methods,  23. 
Inflammation,  109. 
Influenza,  bacillus  of,  118. 
Infusoria,     260;     feeding,     262;      macro- 

and   micro-nucleus    of,    263;     quieting, 

262;   staining,  263;   to  see  cilia  of,  263. 
Injecting  animals  for  preservation,  31. 
Injection:   by  continuous  air-pressure,  86; 

by  syringe,  84,  90;  double,  87;  through 

femoral  artery,  90;    triple,  87. 
Injection  mass,  83;    blue,  83;    cold  fluid 

gelatin,    90;     for    gross    anatomy,    92; 

red,  83;    to  keep,  90;    yellow,  83. 
Injection  methods,  25,  83-92;    Locy's  air 

method,  91. 


Ink:   diamond,  58;  for  glass,  58. 
Insect  larvae,  quieting  aquatic,  272. 
Insects:   delicate,  99;    having  hard  shells, 

99;  to  mount,  99;  transparent  and  soft, 

99. 

Intercellular  bridges,  248. 
Intercolated  disks  in  heart  muscle,  242. 
Interpretation  of  prepared  material,  15. 
Intestinal  absorption  of  fat,  246. 
Intestine,  large,  246;  small,  246. 
Intra-vitam  staining,  146. 
Iodine  for  removing  corrosive  sublimate, 

28. 

Iris,  249. 
Iron-hem  atoxylin:       Heidenhain's,        10; 

method,    51;     use    in    cytology,     139; 

with  other   stains,   52. 
Isaacs,  165. 
Isolation  method,  15,  25,  77,  248. 

Jamming  of  paraffin  sections,  45. 

Janus  green,  145,  146. 

Jelly  of  Wharton,  244. 

Jellyfish,  266. 

Jennings,  259,  269,  271. 

Johnson        122,       215;       paraffin-rubber 

method,    43. 
Joris,  91. 
Julin,  132. 

Keiller's  fluid,  31. 

Kidney:  blood  vessels  of,  257;  cortex 
and  medulla  of,  258;  epithelium  of 
uriniferous  tubules,  258;  glomerulus 
and  its  capsule,  258;  injection  of,  87; 
isolation  of  uriniferous  tubules,  258; 
medullary  rays  of,  258;  nerves  of,  258. 

Killing,  16;    general  rules  for,  27. 

Kincaid,  32. 

Kirkham,  131. 

Kleinenberg's  picro-sulphuric,  217. 

Knower's  injection  method,  88. 

Koch-Ehrlich  gentian  violet,  116. 

Kofoid,  259. 

Kornhauser,  64. 

La  Rue,  267. 
Label,  49. 
Labeling,  58. 
Lachrymal  gland,  249. 
Large  objects,  to  cut  in  paraffin,  44. 
Larvae,  soft,  to  mount,  99. 
Larval  stages  of  trematodes,  267. 
Larynx,  256. 

Lavdowsky's  mixture,  214. 
Lecithin,  tests  for,  148. 
Lee,  37,  73,  223,  276. 
Leech,  272. 

Legs  of  insects,  to  mount,  99. 
Leishman's  Roman owsky's  stain,  110. 
Lens,  249;     capsule    and    epithelium    of, 
249;    fibers,  250. 


Index 


285 


Lenses,  176;   doublets,  180;    triplets,  180. 

Leprosy,  bacillus  of,  117. 

Leptothrix,  115. 

Leucocytes,  111,  241;  ameboid  move- 
ments in,  110;  eosinophylic,  111; 
feeding,  110;  granules  of,  111,  240,  241; 
kinds  of,  111;  large  mpnonuclear,  111; 
poly  nuclear  neutrophylic,  111. 

Lewis,  138,  146,  157. 

Lichtgriin,  227. 

Ligament,  244. 

Ligamentum  nuchae,  244. 

Light  for  microscopical  work,  195. 

Light  green,  20,  227. 

Lillie,  11. 

Line-process,  168. 

Linstaedt,  65. 

Lip,  246. 

Lithography,  171. 

Little,  265. 

Liver  fluke,  100;  injecting,  267. 

Liver,  injection  of,  87. 

Living  cells,  to  study  in  Ambystoma,  143. 

Living  tissue,  staining,  146. 

Locke's  solution,  137. 

Locy's  injection  methods,  91. 

Loeffler's  alkaline  methylen  blue,  116; 
blood-serum,  143. 

Long,  130,  131. 

Lugol's  solution,  148,  227. 

Lung,  256;  blood  vessels  of,  256;  elastic 
tissue  of  alveoli,  256;  epithelium  cf, 
256;  fetal,  256;  injection  of,  87,  91. 

Lymph  capillaries,  242. 

Lymph  glands,  241. 

Lymphatics,  injection  of,  88. 

Lymphocytes,  111. 

Lyons  blue,  10,  50,  227. 

MacCallum's  macerating  fluid,  9,  77,  238. 

McClendon,  14. 

McClung,  1,  14,  18,  22,  31,  40,  101,  152. 

Macerated  tissue,  fixation  of,  80. 

Macerating  fluids,  237,  238. 

Maceration,  15,  25,  77,  78. 

Magenta  S,  224. 

Magnifying  power,  197. 

Malarial  parasite,  111,  241. 

Mallory  and  Wright,  276. 

Mallory's     triple    connective-tissue  stain, 

Mall's  differential  method  for  reticulum, 
79. 

M alone,  75. 

Mammal,  stages  of  maturation,  fertiliza- 
tion, and  segmentation  in,  130. 

Mammary  gland,  257. 

Manipulation    of    the    compound    micro- 
scope, 203. 
Mann,  276. 
Manufacturers,  4. 


Marchi's  method  for  degenerating  nerve 

fibers,  252. 
Mark,  130,  131,  158. 
Marrow,  red,  241. 
Mast,  261. 
Mast-cells,  111,  241. 
Maturation  in  mammals,  130. 
Mayer,  20. 

Mechanical  stage,  197. 
Medulla  oblongata,  252. 
Medullary  sheath,  252. 
Medulla  ted  fibers  of  cord  and  brain,  253, 
Medullated  nerve  fiber,  253. 
Medusae,  266. 
Medusoid  forms,  265,  266. 
Meissner's  corpuscles,  253. 
Membrane  of  Descemet,  250. 
Membranes,  silver-nitrate  method  for,  75. 
Mercuric  crystals,  to  remove,  28. 
Mesothelial  cells,  248. 
Metaelagtin,  96. 

Metallic  substances  for  color  differentia- 
tion, 20,  21,  71. 
Metal  L's  for  imbedding,  45. 
Methods,  general  statement  of,  15. 
Methylated  spirits,  13. 

Methylen  blue,  20,  227;  for  bacteria,  114, 
116;  for  impregnation  of  epithelia,  230; 
for  intra-vitam  staining,  228;  for  nerve 
cells,  230;  for  nerves  and  nerve  termina- 
tions, 228,  230;  for  non-striated  muscle, 
230;  for  ordinary  sections,  230;  immer- 
sion methods  with,  229;  Loeffler's 
alkaline,  116;  polychromatic,  228. 

Methyl  green,  20,  146,  147,  231. 
Methyl  violet,  20,  231. 
Microchemical  tests,  30. 
Micrococci,  115. 
Micrococcus  tetrageneus,  118. 
Micro-dissections,  134,  135. 
Micro-injection  methods,  88,  89. 

Micrometer,  197;  ocular,  198;  screw. 
199;  stage,  197;  step,  199. 

Micron,  199. 

Microscope:  binocular,  189;  compound 
with  parts  named,  181,  182;  demon- 
stration, 192;  dissecting,  188,  193,  194, 
196;  manipulation  of  compound,  203; 
optical  principles  of  the,  175;  path  of 
light  rays  through  the  compound,  184; 
principle  of  compound,  179,  184; 
principle  of  the  simple,  178;  -tube, 
sectional  view  of,  182. 

Microscopes,     representative     compound, 

185,  186. 
Microscopical  material  for  general  zoology, 

259-74. 
Microscopical  terms  and  appliances,  187- 

202. 
Microtome,    4;     automatic   celloidin,    60; 

Bardeen  freezing,  68;   Minot  automatic 

rotary,    39;     oiling    the,    43;     Spencer 

rotary,  39;  well,  35. 

Microtome  knife,  sharpening  the,  43. 


286 


Animal  Micrology 


Milk,  257. 

Milky  or  hazy  staining,  57. 

Miller,  124. 

Minot,  128,  134,  221. 

Minute  dissections,  79;  use  of  clove  oil 
in,  80. 

Mirror,  199- 

Mites,  94. 

Mitochondria,  143,  148;  Benda's  method 
for,  144;  Bensley's  methods  for,  144, 
145;  in  living  cells,  146;  in  tissue 
cultures,  146. 

Mitosis:  in  Ascaris,  136;  in  cells  of 
Ambystoma,  142,  143;  in  eggs  of  white 
fish,  141;  in  grasshoppers,  140;  in 
testis  of  crayfish,  140;  in  testis  of 
Necturus,  141 ;  material  for  demonstrat- 
ing, 139,  140. 

Mollusks,  273. 

Monocystis,  261. 

Mosquito,  to  kill  and  mount,  98. 

Mounting,  22;  general  scheme  of,  26; 
objects  of  general  interest,  93. 

Mounts,  temporary,  35. 

Mouse,  use  in  embryology,  130. 

Mouth,  epithelium  of,  245. 

Mouth-parts  of  insects,  to  mount,  80. 

Muci-carmine,  Mayer's,  231. 

Muci-hematein,  Mayer's,  231. 

Mucin,  231,  232,  246;  stains  for,  148,  231. 

Mucoid  connective  tissue,  244. 

Miiller's  fluid,  210. 

Multiple,  staining,  21. 

Muscae  volitantes,  199. 

Muscle:  areas  of  Conheim,  250;  branched 
striated  fibers,  250;  cardiac,  250;  ends 
of  striated  fibers,  250;  fibrillae  in 
striated,  251;  non-striated,  251;  of 
insects,  97;  Purkinje  fibers,  251;  sar- 
colemma  of,  251;  striated,  251;  to 
tendon,  250. 

Muscle-fibers,  isolation  of,  76. 

Muscular  tissue,  250,  251. 

Mussel:  clearing  entire,  104;  cross- 
section  of  gill,  273. 

Myelin,  252. 

Myelocytes,  111. 


Nail,  257. 

Naphthol,  alpha,  148. 

Naphthylamin  yellow,  240. 

Narcotics  in  killing,  16. 

Necturus,  mitosis  in  testis  of,  141. 

Neisser's  method  for  diagnosing  diph- 
theria, 116. 

Nelson,  103,  104. 

Nematodes,  268. 

Nerve:  plexuses  of  alimentary  canal,  246; 
silver-nitrate  method  for,  74. 

Nerve  cells,  Golgi  method  for  71. 
Nerve-endings,  253 ;  gold-chloride  method 

for,  76. 
Nerve-fiber  bundles,  253. 


Nerve  fibers:  degenerating,  252;  ultra- 
epithelial,  252;  medullated,  253;  non- 
medulla  ted,  253;  stain  for  medullated, 
236. 

Nerve-tissue,  staining  in  toto,  104. 

Nervous  system,  251—54;  of  insect,  to 
mount,  79. 

Neurofibrils,  Cajal's  method  for,  75. 

Neuroglia,  253. 

Neurokeratin,  253. 

Neutral  red,  232. 

Neutrophil  granules,  241. 

Nigrosin,  20. 

Nissl's  granules,  148,  230,  253. 

Nissl's  method  of  staining  basophil 
substance  in  nerve  cells,  230. 

Nitric  acid,  17,  25;  decalcifying  fluid, 
239. 

Nodes  of  Ranvier,  253. 

Normal  fluids,  26,  236. 

Normal  saline,  8,  26,  237. 

Nose,  254. 

Nuclear  stains,  20. 

Numerical  aperture,  199. 

Nymph  of  dragon  fly,  272. 

Objective:  achromatic,  187;  apochro- 
matic,  183,  188;  immersion,  195,  196; 
using  the  immersion,  204. 

Objectives:  lens  systems  of,  180;  rating 
of,  183,  187. 

Ocular:  compensating,  191;  demonstra- 
tion, 192;  Huygenian,  180;  negative, 
180;  positive,  180;  projection,  201; 
searching,  191;  working,  191. 

Oculars,  rating  of,  183,  187. 

Olfactory  cells,  254;  nerve  processes  of,  254. 

Oogenesis,  254. 

Opalina,  261. 

Opaque  mounts,  98. 

Optical  center  of  a  lens,  176. 

Orange  G,  10,  20,  52,  232. 

Orcein,  232. 

Organs  varying  in  density,  70. 

Orientation  of  objects  in  the  imbedding- 
mass,  126. 

Orienting  serial  sections,  126. 

Origanum,  oil  of,  23,  63. 

Orth's  lithium  carmine,  114. 

Osmic  acid,  17,  215;  test  for  fat,  147; 
vapor,  216;  washing  out,  18. 

Osmium-bichromate  mixture  in  Golgi 
method,  73. 

Otoliths,  248. 

Ova,  255. 

Ovary,  255. 

Over-correction,  202. 

Oviduct,  255. 

Oxidase  reaction,  148. 

Oxyphil  granules,  240. 

Pacinian  corpuscles,  253. 
Pancreas,  246;   granules  of,  246. 


Index 


287 


Paper  boxes,  preparing,  37. 
Paracarmine,  Mayer's,  232. 
Paraffin,  12,  23;  baths,  11,  12,  14,  36; 

ribbon  carrier,  40;    to  clean,  42. 
Paraffin  method,  26,  36;    compared  Avith 

celloidin  method,  63;    difficulties  likely 

to  be  encountered  in,  45;    for  delicate 

objects,  53. 

Paraffin-rubber  method  of  Johnson,  43. 
Paraffin  sections:    of  large  objects,  44; 

staining  and  mounting,  48. 
Paramecium,  2*60,  261. 
Parfocal,  200. 
Parotid  gland,  246. 
Pathogenic  bacteria,  118. 
Patton,  127. 
Pearl,  214. 
Peaslee,  100. 
Pectinatella,  269. 
Pedesis,  189. 
Pedicellaria,  269. 
Penetration,  200. 
Penis,  255. 
Peter,  157. 
Peyer's  patches,  246. 
Pfluger's  egg-tubes,  255. 
Phloroglucin  method  of  decalcification,  239. 
Photographing  cellular  structures,  150. 
Photographs,  reproduction  of,  171. 
Photomicrography,  150,  200. 
Pickerel,  mitosis  in  egg  of,  141. 
Picric  acid,  17,  18,  216;    as  a  stain,  233; 

decalcifying  fluid,  239;  washing  out,  18 
Picric-alcohol,  216. 
Picro-acetic,  216;   formalin,  9,  29. 
Picro-carmine,  233. 
Picro-sublimate,  217.   v 
Picro-sulphuric,  217;    for  chick  embryos, 

124. 

Pig,  embryology  of,  133. 
Pipette,  egg,  121. 
Placenta,  255. 
Planaria:    killing  and  mounting,  97;    to 

kill  with  pharynx  protruded,  266. 
Plankton,  263. 
Plasmosomes,  stains  for,  149. 
Platelets,  blood-,  105,  111. 
Plating  celloidin  sections,  65. 
Platino-aceto-osmic  mixture,  217. 
Plumatella,  269. 
Pneumococcus,  118. 
Pointer  ocular,  192. 
Polar  bodies,  131,  135,  136. 
Polariscope,  200. 
Polypoid  forms,  265. 
Potash  clearing  method,  103. 
Preservation,  19,  30, 31 ;  in  paraffin,  38, 42. 
Prickle-cells  of  skin,  257. 
Prostate,  255. 

Protoplasmic    currents,    to    demonstrate, 
151. 


Protozoa,    259-63;     permanent    prepara- 
tions of,  263. 
Purkinje  cells,  253. 
Purkinje  fibers,  242,  251. 
Pyridine-silver  method,  75. 
PyrogaUol,  233. 
Pyroligneous  acid,  233. 
Pyroxylin,  23. 

Quince  jelly  for  Euglena  cultures,  260. 
Quinoline  blue,  222. 

Rabbit,  embryology  of,  131-33. 

Rabl's  picro-sublimate,  217. 

Radula  of  snail,  100. 

Hanson,  75. 

Ranvier's  one-third  alcohol,  238. 

Rat,  embryology  of,  131,  133. 

Rath's  picro-sublimate,  217. 

Ray  of  light,  175. 

Razor,  section,  1,  33. 

Reagents,  preparation  of,  7. 

Reconstruction:      blotting     paper,      157; 

geometrical,  156;  practical  exercise  in, 

155;    reconstruction  methods,    154-58; 

use  of  photography  in,  157 ;  wax,  154, 157. 
Record  cards,  5. 
Rectified  spirits,  13. 
Reduction,  test  for  intra-cellular,  148;   of 

drawings,  168,  172. 
References,  276. 
Refraction,  175. 
Relative  merits  of  paraffin  and  of  celloidin, 

63. 

Remak's  fibers,  253. 
Reproductive  organs,  254-55. 
Resolving  power,  201. 
Resorcin-fuchsin,  234. 
Respiratory  organs,  256. 
Reticular  connective  tissue,  244. 
Reticulum,  Mall's  method  for,  79. 
Retina,  250. 

Rhigolene,  for  freezing,  67,  70. 
Rice,  157. 
Riddle,  147. 
Ringer's  solution,  237. 
Ripart  and  Petit,  liquid  of,  211. 
Robertson,  140. 
Rolling  of  paraffin  sections,  46. 
Ross  board,  168. 
Rotifers,  269. 

Rubbing  sections  off  the  slide,  to  avoid,  55. 
Rubin  S,  224. 
Rules,  general,  5. 

Sabin,  90;     modification    of   the    Spalte- 

holz  method,  102. 
Safranin,  20,  234;   and  gentian  violet,  142 

234;  -gentian-orange  preparations,  142; 

use  in  cytology,  139,  142. 
Salivary    glands:     granules    of,    246;     of 

insect,  to  mount,  80. 


288 


Animal  Micrology 


Sandal-wood  oil,  22. 
Sarcinae,  115. 
Sarcolemma,  251. 
Scales  of  insects,  99. 
Scalpel,  1. 
Scammon,  58,  156. 

Scharlach  R,  235;    stain  for  fat,  148. 
Schule,  146. 

Schultze's  iodized  serum,  237. 
Scissors,  dissecting,  1. 
Sclera,  250. 

Scraping  of  microtome  knife,  46. 
Scratches  across  paraffin  sections,  46. 
Scratching  noise  of  microtome  knife,  45. 
Sealing  bottles,  etc.,  31. 
Sealing  cover-glasses,  95. 
Sebaceous  glands,  257. 
Secretion  antecedents,  149. 
Section  methods,  15,  26. 
Section  razor,  33.     ( 

Sectioning:  by  the  freezing  method,  67; 
injected  organs,  86. 

Sections:  affixing,  23;  cutting  celloidin, 
60;  cutting  frozen,  68;  cutting  paraffin, 
38;  flooding  with  the  dye,  58;  free- 
hand, 33;  preserving  frozen,  33;  wash- 
ing off  of,  57. 

Segmentation  in  mammals,  130. 

Semicircular  canals,  248. 

Seminal  vesicle,  255. 

Seminiferous  tubules,  255. 

Serial  sections,  orienting,  126. 

Sex-chromosomes,  151. 

Shading  drawings,  161,  166. 

Shadows,  in  drawing,  162. 

Sheet  method  for  celloidin  sections,  65. 

Shellac  for  cell-making,  101. 

Shop  skeleton  letters  for  hand  labeling,  164. 

Siedentopf,  202. 

Sihler's  hemotoxylin  method,  104. 

Silver  nitrate,  235;   methods,  74. 

Skin  and  its  appendages,  256-57;  blood 
vessels  of,  257. 

Slide,  box,  1;  marking,  1,  58;  standard,  1. 
Small  intestine,  epithelium  of,  245. 
Small  objects,  transferring,  30. 
Small  pieces,  sectioning  free-hand,  34. 
Smear  preparations:  bacteria,  112;  blood, 
106;    to  show  mitosis  in,  141. 

Smegma  bacilli,  110. 
Smith,  87,  129,  259,  262. 
Snails,  to  kill  expanded,  273. 
Sobotta,  131. 

Sodium-chloride  dissociator,  238. 
Solutions,  rules  regarding,  5. 
Spaeth,  136. 

Spalteholz  method  of  clearing  total 
specimens,  102. 

Spawning  fish,  136. 

Spectrum:  secondary,  183,  188;  tertiary, 
184,  188. 


Spermatogenesis,  255. 

Spermatozoa,  255,  265. 

Spinal  cord,  253. 

Spinal  ganglia,  254. 

Spindles,  140,  149. 

Spirillum,  115;  of  Asiatic  cholera,  118. 

Spleen,  241;    blood  vessels  of,  242. 

SpUtting  of  paraffin  sections,  46. 

Sponges,  264. 

Spores  of  bacteria,  staining,  118. 

Sporozoa,  261. 

Sputum,  to  examine  for  tubercle  bacilli, 

Staining,     19;      and     mounting     paraffin 

sections,  48;  causes  of  failure  in,  56,  57; 

celloidin  sections,   in  hematoxylin  and 

eosin,  61;   in  bulk,  52     progressive,  24; 

regressive,  24. 
Stains,  218-36;    classes  of,  20;    renewing, 

55. 

Staphylococci,  115. 
Staphylococcus     pyogenes:       alba,      118; 

aureus,  118. 
Starfish,  269. 
Stenders,  4. 
Stentor,  259. 
Stephenson,  260. 
Sting,  to  mount,  79. 
Stipple-board,  168. 
Stomach,     247;      cardiac    end    of,     247; 

epithelium    of,    245;     fundus    of,    247; 

pyloric  end  of,  247. 
Stoppers,  to  remove,  14. 
Streeter,  135,  156. 
Streptococci,  115. 
Streptococcus  capsulatus,  118;    pyogenes, 

118. 

Strop  for  microtome  knife,  44. 
Sublingual  glands,  247. 
Subinaxillary  glands,  247. 
Sudan  III,  235;    stain  for  fat,  147. 
Sulphalizarinate  of  soda,  144. 
Supplies  required,  1. 
Supporting  tissues,  242,  245. 
Supporting  vials,  31. 
Suprarenal  glands,  258. 
Sweat  glands,  257. 
Sympathetic  ganglia,  254. 
Synovial  villi,  245. 
Syphilis,  treponema  of,  117. 

Table,  imbedding-,  12. 

Table  of  equivalent  weights  and  measures, 

274. 
Table  of  tissues  and  organs  with  methods 

of  preparation,  240-58. 
Tactile  corpuscles,  254. 
Tactile  menisci,  254. 
Tandler,  90. 
Tapeworms,  100,  267. 
Tap-water  for  washing  sections  stained  in 

hematoxylin,  55. 


Index 


289 


Tashiro's  apparatus  for  measuring  carbon- 
dioxide  output,  153. 

Taste-buds,  247. 

Teasing,  15,  25,  77,  78. 

Tcichmann's  crystals,  106. 

Teleosts,  artificial  fecundation  in,  135; 
care  of  eggs,  136;  embryology  of,  128- 
29;  to  preserve  eggs  of,  129;  to  study 
living  eggs  of,  129. 

Tellyesnicky's  fluid,  209. 

Temperature,  effect  on  paraffin  sectioning, 
43. 

Tendon,  245;    cells,  245;    to  muscle,  245. 

Terminal  bars,  249. 

Testis,  255;  for  avian  and  mammalian 
spermatogenesis,  153. 

Tests  for  certain  cellular  structures,  147,. 
148. 

Tetanus,  bacillus  of,  118. 

Tetracocci,  115. 

Thermometers,  275. 

Thin  sections:  compared  with  thick,  34; 
to  cut,  43;  treatment  of,  55. 

Thionin,  235. 

Thymus  gland,  241,  256. 

Thyroid  gland,  256. 

Tigroid  granules  (substance),  230,  253. 

Tilt  of  microtome  knife,  40. 

Tissues:  cultivation  of  removed,  137; 
length  of  time  required  for  staining,  56 ; 
which  crumble,  43. 

Toisson's  solution,  108. 

Toluidin  blue,  20. 

Toluol,  22. 

Tongue,  247;  papillae  and  folliculi  of,  247. 

Tonsil,  247. 

Tooth,  decalcified,  81,  247;  development 
of,  247;  enamel  prisms  of,  247;  grind- 
ing, 82;  ground,  247;  odontoblasts, 
247;  sectioning  decalcified,  81. 

Trachea,  256. 

Tracheae  of  insects,  Golgi's  method  for,  73. 

Transparent  larvae,  94. 

Trematodes,  266;   larval  stages  of,  267. 

Trichinella,  268;  to  demonstrate  living, 
268. 

Trypan  blue,  146. 

Trypanosoma,  261. 

Tube-length,   181.   182,   184,  201. 

Tubercle  bacilli,  117,  118. 

Turner,  260. 

Turning  cells,  91. 

Turntable,  94. 

Turpentine,  22,  55. 

Typhoid,  bacillus  of,  118 


Ultra-microscopy,  201. 
Umbilicus,  255. 
Under-correction,  202. 
Unna,  230,  232. 
Urea,  use  in  cytology,  152. 


Ureter,  258. 
Urethra,  255,  258. 
Urinary  organs,  257,  258. 
Uterus,  255. 

Vagina,  255. 

Van  Beneden,  132. 

Van  Gehuchten,  218. 

Van  Giesen's  stain,  224. 

Variation  in  thickness  of  paraffin  sections, 

46. 

Vas  deferens,  255. 
Vein,  242. 

Vision,  conventional  distance  of,  183. 
Visual  purple,  in  rods,  250. 
Volvox,  aureus,  261;  globator,  261. 
Von  Ebner's  decalcifying  fluid.  239. 
Vorticella,  261. 

Wagner,  31,  92. 

Waite,  14. 

Walker,  83,  87. 

Walton,  30,  101,  129,  261. 

Washing,  17;  devices,  32. 

Watch-glass,  Syracuse,  4. 

Water  method  for  affixing  sections,  23. 

Water-mites.  94. 

Wax  plates:    anchorage  by  twisted  wires, 

157;   methods  for  rapidly  cutting,  157; 

preparing  for  reconstructions,  154,  155. 
Weber,  156. 
Weigert-Pal   stain  for   medullated   nerve 

fibers,  236. 
Weigert's  borax-ferricyanide,  145;  elastic 

tissue  stain,  234;    method  for  bacteria, 

114. 
Weights  and  measures,  table  of  equivalent, 

274. 

White  blood  corpuscles,  241. 
White  objects,  to  orient,  42. 
Whitman,  87,  121. 

Whole  objects,  mounting,  26,  93,  96,  97. 
Wildman,  144. 

Window  method  for  chick  embryo,  138. 
Wings  of  insects,  98,  99. 
Wintergreen,  oil  of,  22. 
Wood's  metal,  91. 
Worcester's  fluid,  214. 
Working  distance,  202. 
Wright,  202,  276;   stain  for  blood,  110. 
Wrinkles  in  paraffin  sections,  45. 

X-element,  151. 

Xylol,  22,  36;    for  removing  paraffin,  55. 

Zenker's  fluid,  8,  36,  209;   fixing  with,  28; 

formalin  mixtures,  209,  210. 
Ziehl-Neelson  carbol-fuchsin,  116. 
Zoology,  materials  for  a  course  in  generaL 

259-74. 
Zsigmondy,  202. 


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